Looking for yet another additive to try during optimization? Try glycerol. Typical concentrations in the drop can range from 3 to 15%. Glycerol may limit non-specific aggregation and could be useful if you plan to use cryo techniques during data collection.

When adding glycerol to the vapor diffusion crystallization experiment be sure to have a higher glycerol concentration in the reservoir than the drop otherwise you may find that your drops will get biggger and not smaller. Reference: Vera,L., Czarny, B., Georgiadis, D., Dive, V., Stura, E.A. (2011) Practical Use of Glycerol in Protein Crystallization. Cryst. Growth & Des. 11 :2755–2762

When trying to decide which additive might be useful during crystallization, try the following. If one has crystals and wants to try using additives to improve the crystal size or quality go back to the initial crystallization screening plates. Review the plates, looking for conditions where no precipitate nor crystals were/are observed. Now, review the results, looking for a common reagent ingredient present where no precipitant or crystals are found. For example, one might find that all drops with isopropanol remained clear. One could then try adding isopropanol to the current crystallization condition to see if the isopropanol could improve the crystal size and/or quality. If one observes a difference in the crystal in the presence of isopropanol, then one might consider evaluating other additives in the class of alcohols such as ethanol, methanol, tert-butanol and others. If one has no crystals, but plenty of precipitate, phase separation and clear drops, follow the above analysis and try adding the common reagent ingredient found in clear drop to those drops which contained precipitate or phase separation. It is possible this agent could improve or at least manipulate sample solubility. The above tip was submitted from Jarmila Jancarik from the laboratory of Sung Ho Kim at the University of California Berkeley. Thank you Jaru! When using this approach it might be reasonable to discern between concentration independent and dependent precipitation when trying to decide which agent to pursue as an additive. Try evaluating the concentration independent agents first and then look at the other agents if sample quantity permits. For example, if one observe precipitate in 15 to 40% PEG but not in 5 to 10% PEG, it might simply be a concentration. However, if one observes no precipitate in 45 to 60% v/v MPD one could guess that MPD is a reasonable agent to evaluate as an additive.

Although Hampton Research does offer a specifically formulated Additive Screen (HR2-428), here is a tip when one already has a sparse matrix screen or two laying about the lab. When screening additives try adding 50 microliters of each Crystal Screen reagent to 950 microliters the "best" crystallization conditions thus far in order to see if any of the reagents in Crystal Screen might serve as good additives. Crystal Screen 2 is an especially good kit for this technique since Crystal Screen 2 contains numerous divalent cations, Jeffamine® Reagent, and a few other "novel" agents. Jeffamine® is a registered trademark of the Huntsman Petrochemical Corporation Reference Crystallization of foot-and-mouth disease virus 3C protease: surface mutagenesis and a novel crystal-optimization strategy. Acta Crystallogr D Biol Crystallogr. 2005 May;61(Pt 5):646-50. Epub 2005 Apr 20. Birtley JR1, Curry S.

Try benzylkonium chloride (Fluka 12060). This cationic surface active agent has been reported to be useful as a crystallization additive with membrane proteins and may be useful for soluble proteins. We've been using it in the drop between 1 and 3% w/v in water. Try a 10% w/v stock solution in water and dilute into the drop to 1-3%.

The presence of thiosulfate in the protein solution was essential to promote crystal growth and to avoid the formation of unstable and weakly diffracting crystals(1); this is likely to be a consequence of the intrinsic capability of the reduced thiol group of the active-site cysteine to form disulfide bridges, leading to the destabilization of the protein native structure. Sulfane sulfur-donor compounds such as Na2S2O3 are likely to either keep the protein in the persulfurated form or to prevent intermolecular disulfide bridges leading to unfolding and aggregation(2). References 1. Crystallization and preliminary crystallographic characterization of LmACR2, an arsenate/antimonate reductase from Leishmania major. D. Bisacchi, Y. Zhou, B. P. Rosen, R. Mukhopadhyay and D. Bordo. Acta Cryst. (2006). F62, 976-979. 2. Bordo, D., Forlani, F., Spallarossa, A., Colnaghi, R., Carpen, A., Bolognesi, M. & Pagani, S. (2001). Biol. Chem. 382, 1245–1252.

Try dissolving the small molecule additive into paraffin or silicon oil, and use this mixture to cover the sample drop. This can be used with sitting drop vapor diffusion or with microbatch under oil. The oil acts as a reservoir that may contain excess small molecule that (you hope) will be fed into the crystals.

Try 3 to 5% 1,2-propanediol in the protein buffer as a substitute for glycerol to stabilize proteins. Annie Hassell, Glaxo Smithkline 2009

Non-specific aggregation problems? Watch your sample buffer. The selection of an appropriate sample buffer/pH can often ward off aggregation problems.

Aggregation can be a deterrent to the crystallization of biological macromolecules including proteins, peptides, and nucleic acids. The presence of sample aggregation can be detected by either dynamic light scattering or native gel electrophoresis. Aggregation might be caused by hydrophobic patches on the surface of the sample, differently charged isoforms, differently phosphorylated isoforms, mixtures of methylated and non-methylated samples, glycosylation, as well as electrostatic interactions. Aggregation can be due to autologous aggregation where the protein is aggregating with itself or heterologous contamination where the sample is aggregating with other proteins. In the case of heterologous contamination, further purification of the sample should be seriously considered. In the case of autologous aggregation that precludes crystallization one might consider: 

Using molecular biology to manipulate intra and inter molecule interactions by modifying the sample sequence (alter, add, or delete residues). 

Use chemical additives to manipulate sample-sample and sample solvent interactions. 
Detergents 
Chaotropes (urea, guanidine hydrochloride, hydrochloric acid, etc) 
Electrostatic agents 
Alcohols (isopropanol, methanol, ethanol, etc) 
Salts (sodium chloride, potassium chloride, sodium fluoride, etc) 
Polyols (glycerol, PEG 400, etc) 
Ligands, inhibitors, co-factors, and metals 
Use temperature to prevent aggregation (0°C and 60°C) 
Consider a fusion protein 
Remove C-terminus or N-terminus 
Truncate domains 
Remove His-tag 

In some cases aggregates can be removed by centrifugation or filtration. 

In some cases the aggregates can be removed by mixing the sample with the crystallization reagent, allowing the sample to incubate for 15 minutes, centrifuging the sample/reagent mixture, removing the precipitate and setting the drop with the supernatant.

We've had good success using the well solution directly as the foundation of a cryobuffer in several situations where crystals cannot be grown directly in the presence of cryoprotectant, and where crystals don't tolerate transfer to artificial mother liquors. The basic protocol is as follows:

1. Remove 100 microliters of the well solution after crystals have grown

2. Split this sample into two 50 microliter aliquots.

3. Add 7.5 mg of dextrose (glucose) to the first aliquot and 15 mg of dextrose to the second. Dissolve by gentle pipeting with a wide-bore tip. This will give two sequential well solutions that now contain 15% and 30% w/v dextrose. If all the dextrose won't go into the second aliquot, spin hard and remove the supernatant.

4. Transfer the crystal to aliquot 1, equilibrate for 3 minutes, then to aliquot number 2, then freeze. We've had a few crystals that routinely crack or blow up when transfered to artificial mother liquor that behave well when transfered to well solution plus glucose. We assume that there is some aspect of the crystal drop (pH, ionic tension, precipitant concentration) that is more effectively reproduced within the well than by separately prepared mother liquors. The nice thing about the protocol above is that you don't get much of a volume increase when dry dextrose is dissolved in the well solution, so the components in the solution are not diluted.

Finally, if you don't get a really good freeze, you can try to add about 5% v/v glycerol to aliquot 2 in addition to the 30% w/v dextrose. Reference: Personal communication from Barry Stoddard, Fred Hutchinson Cancer Research Center

High molecular PEGS are also good cryoprotectants. If crystals are obtained from relatively high concentration of PEGS (e.g., 30% of PEG 3350), you can cryoprotect them simply by raising the concentration of the PEG a little bit (e.g., 40% of PEG 3350). 

J. Appl. Cryst. (2006). 39, 244-251 
Effects of cryoprotectant concentration and cooling rate on vitrification of aqueous solutions. V. Berejnov, N. S. Husseini, O. A. Alsaied and R. E. Thorne 
Synopsis: Critical concentrations required for vitrification of aqueous solutions are determined for fourteen common cryoprotectants, for sample volumes ranging over four orders of magnitude and covering the range of interest in protein crystallography.

To mount very thin crystals onto cryoloops, first dip the nylon loop into 0.5% Formvar solution (Fluka # 09819) to form a thin film. The film provides extra support for fragile crystals, and can result in much sharper reflections with just slightly higher background. To clean the loop, dip it in alcohol to dissolve the support. Two notes: (1) the technique works only for crystals grown without organic solvents, and (2) take precautions not to breathe vapor from the formvar solution--the solvent is 1,2 dichloroethane. Formvar is a standard support for electron microscopy grids.

Reduced radiation damage. 

Decreased thermal motion and disorder. 

Potential for improved resolution. 

Increased crystal lifetime. 

Crystals can be stored and shipped.

Cryo Trouble? Give the following tips a try next time your crystal is fussy about freezing. 

Try X-ray data collection at room temperature. 

Evaluate other cryoprotectants. Try CryoPro from Hampton Research, which contains 36 unique cryopreservation reagents. 
For more information visit http://hamptonresearch.com/product_detail.aspx?cid=30&sid=189&pid=30 

Mixing of different cryoprotectants can have a superior protective effect over single component cryoprotectants of the same total concentration. 

Change the rate of cooling. 

a) Accelerate the rate of cooling. The fastest cooling rates have been achieved by blowing off the gas layer on liquid nitrogen during plunge cooling. (Hyperquenching for protein cryocrystallography, M. Warkentin et al, J. Appl. Cryst. (2006). 39, 805-811) 

b) Slow the rate of cooling. Perhaps better suited to crystals with smaller solvent channels. The key to successful slow cooling of protein crystals is to carefully and completely remove all of the solvent from the surface of the crystal using oil such as Paratone-N, Perfluoropolyether, Mineral, Silicon, NVH or other. Remove ALL of the liquid from the surface of the crystals when using oil. (Slow cooling and temperature-controlled protein crystallography, Warkentin et al 10.1007/s10969-009-9074-y and Slow cooling of protein crystals, Warkentin et al, Volume 42, Part 5, Pages 944-952, October 2009) For more information visit http://hamptonresearch.com/menus.aspx?id=3&sid=138 

Using salt as a crystallization reagent? Many salts are cryosalts, including malonate, formate, citrate, tartrate, acetate, Tacsimate and other organic acids, ammonium sulfate (>3.5 M), lithium sulfate, lithium chloride and other alkylammonium salts. For cryo try increasing your salt concentration by 20%. (Cryosalts: suppression of ice formation in macromolecular crystallography, K. A. Rubinson et al, Acta Cryst. (2000). D56, 996-1001 doi:10.1107/S0907444900007587. 

Try the identical cryo procedure again with another crystal. 

Vary the time and temperature of the crystal handling steps. 

Check the liquid nitrogen level in your dewar and maintain a consistent level, day to day, week to week, month to month, year to year. 

Try annealing. (Macromolecular crystal annealing: Techniques and cases studies. Bunick et al, The Rigaku Journal Vol. 15/ number 2/ 1998 and Macromolecular crystal annealing: overcoming increased mosaicity associated with cryocrystallography, Harp et al. (1998). Acta Cryst. D54, 622-8 

Match the osmotic pressure of your cryoprotectant to the osmotic pressure of the reagent producing the crystal. Crystallization reagents with lower salt concentrations require a higher percentage of cryoprotectant for cryo protection than crystallization reagents with higher salt solutions (Cool data: quantity and quality. Elspeth Garman. Acta Cryst. (1999). D55, 1641-1653.). Osmolality tables (Weast, R. C. (1988-1989). Editor. Handbook of Chemistry; Physics, 69th ed. Boca Raton, Florida: CRC Press) can be used to estimate the osmolality of reagents. Another trick is to slowly concentrate a drop of the mother liquor by leaving the drop open to air and allowing the drop to slowly dry down, checking the mother liquor for clear glass freeze every few minutes. If you do not have X-rays to check for clear glass freeze, you can guesstimate by carefully placing the dewar under a dissecting microscope with overhead lighting. Focus on the surface of the liquid nitrogen, and bring the cooled loop into view just above the surface of the liquid nitrogen, where it is cold enough for guesstimate freezing. 

Try high pressure cooling. (High-pressure cooling of protein crystals without cryoprotectants, Kim et al, Acta Cryst. (2005). D61, 881–890) 

Cryoprotection of delicate crystals - Artem Evdokimov's humble recipe. 
http://www.xtals.org/crystal_cryo.pdf 

Two words. Liquid propane. 
http://cars9.uchicago.edu/biocars/pages/flashcooling.shtml 

Roger S. Rowlett's Cryoprotectant Strategy 
1. Transfer crystals to mother liquor plus 30%v/v glycerol or ethylene glycol (sometimes lower depending on crystallization 
reagent). 
2. Transfer crystals to mother liquor plus 30%w/v glucose (or try sequential soaks in mother liquor plus 15%w/v and then 30%w/v glucose. Just a few seconds or minutes is usually enough). Glucose or other sugars often work when glycerol or ethylene glycol fails. 
3. Try the "no-fail" in situ cryo method, which is a gradual buildup of cryoprotectant. This method is especially appropriate for crystals that cannot tolerate direct transfer to cryoprotectant solution, or for crystals that are especially sensitive to concentration changes in the mother liquor driven by drop evaporation. In our laboratory this method is routinely used with success on otherwise very sensitive crystals. This particular method is adapted for hanging drop crystallization. Ligands can be soaked in at the same time as cryopreservation if included in the cryoprotectant solution at 125% of the final, desired concentration. 
a) Prepare a solution of artificial mother liquor plus 30% w/v glucose (40% v/v glycerol or another cryoprotectant can be substituted) 
b) Remove a coverslip containing a drop with crystals to be cryoprotected and add 0.25 drop volume (DV) of cryoprotectant solutions (e.g. for a 4 uL drop add 1 uL of cryoprotectant solution). Replace coverslip on well and let stand for 5 minutes. Examine the crystals for cracking and/or dissolution. 
c) Repeat the previous step with the following additional cryoprotectant additions: 0.25 DV, 0.50 DV, 1.00 DV, 2.00 DV. After each addition replace the coverslip over the well and let stand for 5 minutes. Examine crystals for cracking and/or dissolution. 
d) After the last addition and 5 minute incubation, remove coverlip, fish out crystals with mounting loops and freeze directly in liquid nitrogen. The final glucose concentration will be 24%, sufficient to protect most crystallization solutions from ice formation upon freezing in liquid nitrogen. 
This is very gentle, and often works when #1 and #2 does not, but in our hands nearly always increases mosaicity. (But mosaic is better than no diffraction.) 
4. Try dragging crystals though Paratone-N to remove surface water from the crystal. This actually nearly always works for us, but is more fussy than #1 or #2, and it is easier to damage crystals during manipulation because of the viscosity of the oil. 
I normally plunge protected crystals into liquid nitrogen after mounting. 
Ice rings are a good indication of poor cryoprotection, but lack of diffraction could just be your crystals, too. For our latest dataset, we just sorted through 38 crystals until we found a good one. The key, as it turned out, is that all of our beautiful large crystals were apparently difficult to visualize disordered stacks of plates (we didn't notice this until some fractured during cryo-soaks) whereas some of the small crystals were actually single crystals. We selected a decently diffracting small one and took loooooooooooong frames to get a good data set. (Roger S. Rowlett, Professor, Colgate University Presidential Scholar, Department of Chemistry, Colgate University, NY, USA) 

Dig around in the cryoprotectant database for ideas. http://idb.exst.jaxa.jp/db_data/protein/search-e.php 

Try 10% glycerol. (Jim Pflugrath) 

Try 10 to 20% 2,3-butanediol. It can also reduce mosaicity. Try a quick dip, 30 to 60 seconds. 

Fine tune the cryo concentration by screening 2.5% concentration increments, dipping a loop in the cryo buffer, freezing it and collecting X-Ray diffraction data to find the minimum concentration required that produces a clear glass and no ice rings. 

Try dragging the crystal through a 1:1 mix of Paratone-N and Mineral oil until most or all the mother liquor from surrounding the crystal has been removed. (David Briggs) 

Look back in your screen plates for a different crystal in different reagent, do not pass go, do not collect $200.00 and start over at the top of this tip. 

(Be aware of the potential for) detergent concentration mismatch between your mother liquor and the cryosolution. This particularly happens with vapor diffusion setups: there is a delicate balance of "free" detergent in the mother liquor versus the proportion of the detergent which is bound to the protein. Dropping a xtal into the cryosolution shocks the crystal with a bolus of extra free detergent. Hence, and counterintuitively, you may need to reduce the detergent concentration in the cryosolution to keep everything in balance. Try titrating down from 1% to even as low as 0.4% in the cryosolution. Under the conditions you are using the CMC of bOG is suppressed below the usual 0.67% (w/v). 

Also, the behavior of many of the alkyl glycoside detergents is very temperature sensitive. So be careful about the temperature of all the solutions you use. 

R. Michael Garavito, Ph.D. 
Submitted to CCP4 bulletin board February 2007 
Edited by Hampton Research Corp. 

Soak the CryoLoop in a 0.5% detergent solution in deionized water for 15 minutes. Gently rinse the CryoLoop in deionized water. Allow the CryoLoop to air dry. 

If more rigorous cleaning is desired or one does not wish to risk the presence of trace amounts of residual detergent, soak the CryoLoop in 5.0 M Urea formulated in deionized water for 15 minutes. Gently rinse the CryoLoop in deionized water. Repeat three times. Allow the CryoLoop to air dry.

Increase the concentration of ammonium sulfate (cryosalt) (or primary salt acting as a precipitant) in increments of 10% in the presence of 5 to 10% v/v Glycerol. 

Evaluate 25-30% w/v glucose, or trehalose or sucrose as cryoprotectants for crystals grown in Ammonium sulfate. Acta Cryst. (2002). D58, 1664-1669. Crystallization of RNA/protein complexes. M. Garber, G. Gongadze, V. Meshcheryakov, O. Nikonov, A. Nikulin, A. Perederina, W. Piendl, A. Serganov and S. Tishchenko. 

15 to 30% v/v Ethylene glycol, DMSO, or Glycerol. 

Try 1 – 2 M Sodium malonate as a cryoprotectant. Acta Cryst. (2003). D59, 2356-2358. Malonate: a versatile cryoprotectant and stabilizing solution for salt-grown macromolecular crystals. T. Holyoak, T. D. Fenn, M. A. Wilson, A. G. Moulin, D. Ringe and G. A. Petsko. 

Place a drop of 75% v/v Paratone-N, 25% v/v Paraffin Oil next to the drop containing the crystal. Remove the crystal from the drop using a CryoLoop and dip the crystal into the oil. Keeping the CryoLoop and crystal immersed, gently move the CryoLoop containing the crystal from the oil into the reagent, back into the oil, back into the reagent, repeating this several times to remove some of the aqueous (reagent) layer from the crystal surface. Cryo cool the crystal. 

Got ice rings and want to get rid of them? 

While the crystal is in the beam, carefully try the following. 

Try annealing the crystal by blocking the stream for a few seconds, allowing the crystal to thaw, then unblock the stream to cool the crystal again. (Reference: New techniques in macromolecular cryocrystallography: macromolecular crystal annealing and cryogenic helium, B. Leif Hansona, Constance A. Schallb and Gerard J. Bunick, Journal of Structural Biology 
Volume 142, Issue 1, April 2003, Pages 77-87) 

Try washing the crystal with liquid nitrogen. Using a pipette, carefully aspirate liquid nitrogen into the pipette, then dispense several drops or even a gentle stream over the crystal and mount. A liquid nitrogen wash can also be used to remove creeping ice from caps, pins and loops.

Removal 
With one hand, using pliers, grasp the Mounted CryoLoop. Using your the other hand, hold the CrystalCap between your thumb and finger(s). With gentle authority twist the Mounted CryoLoop and Cap in opposing directions and pull the Mounted CryoLoop from the CrystalCap. 

Replacement 
Select a Mounted CryoLoop with the desired CryoLoop size. Notice the notches on the Mounted CryoLoop. A standard length Mounted CryoLoop is created by snapping the MicroTube at the second notch from the bottom. Remove the end using your fingers or diagonal cutter. Apply a drop of Super Glue to the top opening of the CrystalCap. Insert the Mounted CryoLoop into this opening and push until the Mounted CryoLoop is stopped by the bottom of the CrystalCap. Epoxy may be used instead of Super Glue for a more permanent fixation of the Mounted CryoLoop to the CrystalCap. 

Using salt as a crystallization reagent?

Many salts are cryosalts, including malonate, formate, citrate, tartrate, acetate, Tacsimate and other organic acids, ammonium sulfate (>3.5 M), lithium sulfate, lithium chloride and other alkylammonium salts. For cryo try increasing your salt concentration by 20%. (Cryosalts: suppression of ice formation in macromolecular crystallography, K. A. Rubinson et al, Acta Cryst. (2000). D56, 996-1001 doi:10.1107/S0907444900007587. 

Moisture accumulcation in cryogenic dry shipper dewars can lead to frost forming about mounted crystals stored in pucks. One can rinse the frost away using liquid nitrogen. But to avoid frost formation in the fist place, when not in use, be sure to store cryogenic dry shipper dewars inverted to remove moisture and to flush the dewar with dry air to remove moisture. 

Another option is to keep the cryogenic dry shipper filled with liquid nitrogen at all times. This can often be accomplished with weekly checks and fills. 

A dry liquid nitrogen supply can also minimize ice accumulation. 

Practical macromolecular cryocrystallography by J.W. Pflugrath

Acta Crystallogr F Struct Biol Commun. 2015 Jun 1; 71(Pt 6): 622–642. Published online 2015 May 27. 

Abstract 
Cryocrystallography is an indispensable technique that is routinely used for single-crystal X-ray diffraction data collection at temperatures near 100 K, where radiation damage is mitigated. Modern procedures and tools to cryoprotect and rapidly cool macromolecular crystals with a significant solvent fraction to below the glass-transition phase of water are reviewed. Reagents and methods to help prevent the stresses that damage crystals when flash-cooling are described. A method of using isopentane to assess whether cryogenic temperatures have been preserved when dismounting screened crystals is also presented. 

Acta Cryst. (2009). D65, 823-831 [ doi:10.1107/S0907444909017958 ] Characterization of gadolinium complexes for SAD phasing in macromolecular crystallography: application to CbpF. R. Molina, M. Stelter, R. Kahn and J. A. Hermoso. Synopsis: In SAD phasing, the efficacy of cocrystallization versus soaking in the binding of the different Gd complexes, their mode of interaction and a comparative study using synchrotron radiation and rotating-anode generator, were tested. 

Novel sample preparation technique for protein crystal X-ray crystallographic analysis combining microfluidics and acoustic manipulation 
S. Oberti, D. Möller, S. Gutmann, A. Neild and J. Dual. J. Appl. Cryst. (2009). 42. Synopsis: A technique for sample preparation prior to X-ray crystallographic analysis of protein crystals involving the use of acoustic radiation forces and a micro-machined fluidic device is introduced. Crystals can be positioned in a sequence along a channel and then individually removed with a loop, which is then mounted onto a goniometer. 

Successful protein cryocooling for powder diffraction. M. J. Jenner, J. P. Wright, I. Margiolaki and A. N. Fitch. J. Appl. Cryst. (2007). 40, 121-124 

What can be done with a good crystal and an accurate beamline? Jiawei Wang, Miroslawa Dautera and Zbigniew Dauter. Acta Cryst. (2006). D62, 1475–1483. 

Automation of sample mounting for macromolecular crystallography. F. Cipriani et al. Acta Cryst. (2006). D62, 1251-1259. 

How to avoid premature decay of your macromolecular crystal: A quick soak for a long life. B. Kauffmann, M.S. Weiss, V.S. Lamzin, and A. Schmidt. Structure 14, 1099-1105, July 2006. 

Using image analysis for automated crystal positioning in a synchrotron X-ray beam for high-throughput macromolecular crystallography. Andrey et al. J. Appl. Cryst. (2004). 37, 265-269. 

Use of dry paraffin oil and Panjelly in the xenon derivatization of protein crystals. J. Appl. Cryst. (2002). 35, 117-119. 

CATS: a cryogenic automated transfer system installed on the beamline FIP at ESRF. Ohana et al. J. Appl. Cryst. (2004). 37, 72-77. 

Applications of the streak seeding technique in protein crystallization. Stura and Wilson. Journal of CrystaL Growth, 110 (1991) 270-282. 

X-ray scattering studies of Aspergillus flavus urate oxidase: towards a better understanding of PEG effects on the crystallization of large proteins. D. Vivares and F. Bonnete. Acta Cryst. (2002). D58, 472-479. 

Flash colling and annealing of protein crystals. S. Kriminski et al. Acta Cryst. (2002). D58, 459-471. 

Resolution improvement from 'in situ annealing' of copper nitrite reductase crystals. Mark J. Ellis et al. Acta Cryst. (2002). D58, 456-458. 

Phasing possibilities using different wavelength with a xenon derivative. Santosh Panjikar and Paul Tucker. J. Appl. Cryst. (2002). 35, 261-266. 

Automation of the EMBL Hamburg protein crystallography beamline BW7B. Ehmke Pohl et al. J. Synchrotron Rad. (2004). 11, 372-377. 

Protein crystallography in a vapour stream: data collection, reaction initiation and intermediate trapping in named hydrate protein crystals. Sjogren, T. et al. J. Appl. Cryst. (2002). 35, 113-116. 

Radiation driven collapse of protein crystals. Ian K. Robinson et al. J. Synchrotron Rad. (2006). 13, 1-7. 

Beam-size effects in radiation damage in insulin and thaumatin crystals.C. Schulze-Briese et al. J. Synchrotron Rad. (2005) 12, 261-267. 

Will reduced radiation damage occur with very small crystals? Colin Nave et al. J. Synchrotron Rad. (2005) 12, 299-303. 

Towards an understanding of radiation damage in cryocooled macromolecular crystals. Colin Nave et al. J. Synchrotron Rad. (2005) 12, 257-260. 

A review of techniques for maximizing diffraction from a protein crystal in stilla. Janet Newman. Acta Cryst. (2006) D62, 27-31. 

Measurement of the density, Composition and Related Unit Cell Dimension of some Protein Crystals. F.M. Richards, et al. Journal of the American Chemical Society, 1954, Volume 76, Iss. 9/2211-2518. 

In situ data collection and structure refinement from microcapillary protein crystallization. Raymond C. Stevens et al. J. Appl. Cryst. (2005) 38, 900-905. 

Protein crystallography microdiffraction. Christian Riekel et al. Current Opinion in Structural Biology 2005, 15:556-562. 

Advances in kinetic protein crystallography. Dominique Bourgeois et al. Current Opinion in Structural Biology 2005, 15:538-547. 

Protein nanocrystallography: growth mechanism and atomic structure of crystals induced by nanotemplates. J. Synchrotron Rad. (2005). 12, 772-778. 

Combining X-ray crystallography and electron microscopy. Michael G. Rossmann et al. Structure, Vol. 13, 355-362, March 2005. 

A new method for predetermining the diffraction quality of protein crystals: using SOAP as a selection tool. Robin Leslie Owen and Elspeth Garman. Acta Cryst. (2005). D61, 130-140. 

Finding a cold needle in a warm haystack: infrared imaging applied to locating cryocooled crystals in loops. Edward.

A method for screening the temperature dependence of three-dimensional crystal formation. Ben Hankamer et al. Acta Cryst. (2006). D62, 559-562. 

Studies of insulin crystals at low temperatures: Effects on mosaic character and radiation sensitivity. B.W. Low et al. Proc Natl Acad Sci U S A. 1966 December; 56(6): 1746–1750. 

Multiwell microbatch crystallization on a thermal gradient. Ronald W. Rousseau et al. Crystal Growth & Design, Vol. 5, No. 4, 2005. 

High-throughput screening: designer screens. Nathan Blow. Nature Methods 6, 105 - 108 (2009). 

The role of bias in crystallization experiments in automated microseeding. Edwin Pozharski et al. Acta Cryst. (2008). D64 1222-1227. 

Assessment of a preliminary solubility screen to improve crystallization trials: uncoupling crystal condition searches. Aude Izaac, COnstance A. Schall and Timothy C. Mueser. Acta Cryst. (2006). D62, 833-842. 

An attempt to increase the efficiency of protein crystal screening: a simplified screen and experiments. Wei Gao, Shi-xing Li and Ru-chang Bi. Acta Cryst. (2005). D61, 776-779. 

Crystallization screens. Compatibility with the lipidic cubic phases for in meso crystallization of membrane proteins. Cherezov and Caffrey. Biophys. J. Vol 81, 2001. 

Rapid crystallization of chemically synthesixed hammerhead RNAs using a double screening procedure. William G. Scott et al. J. Mol. Biol. (1995) 250, 327-332. 

Tapping the protein data bank for crystallization information. Thomas S. Peat et al. Acta Cryst. (2005). D61, 1662-1669. 

Rapid preparation of custom grid screens for crystal growth and optimization. Anne B. Senger and Timothy C. Mueser. J. Appl. Cryst. (2005). 38, 847-850. 

Crystallization optimum solubility screening: using crystallization results to identify the optimal buffer for protein crystal formation. Bernard Collins, Raymond C. Stevens and Rebecca Page. Acta Cryst. (2005). F61, 1035-1038. 

Statistal experimental design of protein crystallization screening revisited. Anita Lewitt-Bentley et al. Acta Cryst. (2004). D60, 1562-1568. 

Protein isoelectric point as a predictor for increased crystallization screening efficiency. Katherine A. Kantardjieff and Barnhard Rupp. Bioinformatics, Vol. 20 no. 14 2004, pages 2162-2168. 

Towards rationalization of crystallization screening for small- to medium-sixed academic laboratories; the PACT/JCSG+ strategy. Anastassis Perrakis et al. Acta Cryst. (2005). D61, 1426-1431. 

Expanding screening space through the use of alternative reservoirs in vapor-diffusion experiments. Janet Newman. Acta Cryst. (2005). D61, 490-493. 

Enhancing protein crystallization through precipitant synergy. Peter D. Kwong et al. Structure, Vol. 11, 1-20, September 2003. 

Shotgun crystallization strategy for structural genomics: an optimized two-tiered crystallization screen against the THermotoga maritima proteome. Page et al. Acta Cryst. (2103). D59, 1028-1037. 

Data mining crystallization databases: Knowledge-based approaches to optimize protein crystal screens. M.S. Kimber et al. Proteins: Structure, Function, and Genetics 51:562-568 (2003). 

Two approaches to the rapid screening of crystallization conditions. Alexander McPherson. Journal of Crystal Growth 122 (1992) 161-167. (Screening reagents and crystallization by isoelectric focusing IEF) 

Using sample techniques in protein crystallization. H. Sheng, W.C. Stallings, A.M. Stevens and R.A. Stegeman. Acta Cryst. (1995), D51, 305-310. 

A survey of protein-protein complex crystallizations. Radaev et al. Acta Cryst. (2006). D62, 605-612.

A size filtration approach to purify low affinity complexes for crystallization. Ignate et al. Journal of Structural Biology, Volume 159, Issue 1, July 2007, Pages 154-157. 

Cross-linking of protein crystals as an aid in the generation of binary protein-ligand crystal complexes, exemplified by the human PDE10a-papaverine structure. O. A. Andersen, D. L. Schönfeld, I. Toogood-Johnson, B. Felicetti, C. Albrecht, T. Fryatt, M. Whittaker, D. Hallett and J. Barker 

Rigorous filtration for protein crystallization. N. E. Chayen. J. Appl. Cryst. (2009). 42. Synopsis: The effect of filtration on the number, size and quality of protein crystals is described. 

Rational mutagenesis to support structure-based drug design: MAPKAP kinase 2 as a case study. Maria A Argiriadi et al. BMC Structural Biology 2009, 9:16. 

A measure of floppiness. Irene Kaganman. Nature Methods 6, 114 - 115 (2009). 

Influence of micro-impurity on protein crystal growth studied by the etch figure method. Guoliang Daia, Xingyu Liua, Gen Sazakib and Xiaogang Zhang. Journal of Crystal Growth, Volume 311, Issue 3, 15 January 2009, Pages 548-552. 

Thioredoxin as a fusion tag for carrier-driven crystallization. Lorenzo Corsini et al. Protein Science (2008), 17:2070-2079. 

Toward rational protein crystallization: A Web server for the design of crystallizable protein variants. Lukasz Goldschmidt, David R. Cooper, Zygmunt S. Derewenda, and David Eisenberg. Protein Science (2007), 16:1569-1576. 

Cleavage of recombinant proteins at poly-His sequences by Co(II) and Cu(II). Martina Andberg, Jussi Jäntti1, Sara Heilimo, Päivi Pihkala, Arja Paananen, Ari M.P. Koskinen, Hans Söderlund, and Markus B. Linder. Protein Science (2007), 16:1751-1761 

‘Crystal lattice engineering,’ an approach to engineer protein crystal contacts by creating intermolecular symmetry: Crystallization and structure determination of a mutant human RNase 1 with a hydrophobic interface of leucines 
Hidenori Yamada, Taro Tamada, Megumi Kosaka, Kohei Miyata, Shinya Fujiki, Masaru Tano, Masayuki Moriya, Mamoru Yamanishi, Eijiro Honjo, Hiroko Tada, Takeshi Ino, Hiroshi Yamaguchi, Junichiro Futami, Masaharu Seno, Takashi Nomoto, Tomoko Hirata, Motonobu Yoshimura, and Ryota Kuroki 
Protein Sci 2007 16: 1389-1397. 

A simple method for improving protein solubility and long-term stability. Alexander P. Golovanov, Guillaume M. Hautbergue, Stuart A. Wilson, and Lu-Yun Lian. J. Am. Chem. Soc. 2004, 126, 8933-8939. (adding 50 mM L-Arg and L-Glu) 

Highly efficient selenomethionine labeling of recombinant proteins produced in mammalian cells. W.A. Barton, D. Tzvetkova-Robev, H. Erdjument-Bromage, P. Tempst, and D.B. Nikolov. Protein Science (2006), 15:2008-2013. 

Fluorescence-Detection Size-Exclusion Chromatography for Precrystallization Screening of Integral Membrane Proteins. Eric Gouaux et al. Structure 14, 673-681, April 2006. 

Rapid and simple protein-stability screens: application to membrane proteins. H.B. Stowell et al. Acta Cryst. (2006). D62, 451-457. 

An automated in vitro protein folding screen applied to a human dynactin subunit. Scheich et al. Protein Science, (2004), 13:370-380. 

Coupling of MBP fusion protein cleavage with sparse matrix crystallizatin screens to overcome problematic protein solubility. Joseph E. Wedekind et al. BioTechniques, Vol. 39, No.4 (2005). 

Glycine residues appear to be evolutionarily conserved for their ability to inhibit aggregation. Fabrizio Chiti et al. Structure, Vol. 13, 1143-1151, August 2005. 

A straight-forward method of optimising protein solubility for NMR. Peter W.A. Howe. Journal of Biomolecular NMR 30: 283-286, 2004. 

Relationship between preferntial interaction of a protein in an aqueous mixed solvent and its solubility. Ivan L. Shulgin and Eli Rickenstein. Biophysical Chemistry, 118 (2005) 128-134. 

A protein molecule in an aqueous mixed solvent: fluctuation theory outlook. Ivan L. Shulgin and Eli Rickenstein. The Journal of Chemical Physics 123, 054909 (2005). 

A preliminary solubility screen used to improve crystallization trials: crystallization and preliminary X-ray strcture determination of Aeropyrum pernix flap endonuclease-1. Tomothy C. Mueser et al. Acta Cryst. (2004). D60, 1674-1678. 

Optimum solubility (OS) screening: an efficient method to optimize buffer conditions for homogeniety and crystallization of proteins. Jarmila Jancarik, Ramona Pufan, Connie Hong, Sung-Hou Kim and Rosalind Kim. Acta Cryst. (2004). D60, 1670-1673. 

On the use of DXMS to produce more crystallizable proteins: Structures of the T. maritima proteins TM0160 and TM1171. Scott A. Lesley et al. Protein Science (2004), 13:3187-3199. 

A rapidly filled capillary mount for both dry powder and polycrystalline slurry samples. R. B. Von Dreele. J. Appl. Cryst. (2006). 39, 124-126. 

Solubility engineering and crystallization of apolipoprotein D. Arne Skerra et al. Protein Science (2006), 15:190-199.

Proteolytic fragmentation of polypeptide release factor 1 of Thermus thermophilus and crystallization of stable fragments. Tin et al. Biochimie 82 (2000) 765-772. 

A sparse matrix approach to the solubilization of overexpressed proteins. Lindwall et al. Protein Engine.

Crystallization of soluble proteins in vapor diffusion for x-ray crystallography. Manuela Benvenuti and Stefano Mangani. Nature Protocols Volume 2 Number 7 2007 1633-1651. 

Crystallization of protein-ligand complexes. 
A. M. Hassell, G. An, R. K. Bledsoe, J. M. Bynum, H. L. Carter, S.-J.J. Deng, R. T. Gampe, T. E. Grisard, K. P. Madauss, R. T. Nolte, W. J. Rocque, L. Wang, K. L. Weaver, S. P. Williams, G. B. Wisely, R. Xu and L. M. Shewchuk. Acta Cryst. (2007). D63, 72-79. 

Physiological aspects: Part 1 in a series of papers devoted to surfactants in microbiology 
and biotechnology. Jonathan D. Van Hamme, Ajay Singh, Owen P. Ward. Biotechnology Advances 24 (2006) 604–620. 

SPINE high-throughput crystallization, crystal imaging and recognition techniques: current state, performance analysis, new technologies and future aspects. Ian M. Berry, O. Dym, R.M. Ensouf, K. Harlos, R. Meged, A. Perrakis, J.L. Sussman, T.S. Walter, J. Wilson, and Albrecht Messerschmidt. Acta Cryst. (2006). D62, 1137-1149. 

Automation of sample mounting for macromolecular crystallography. F. Cipriani et al. Acta Cryst. (2006). D62, 1251-1259. 

A brief history of protein crystal growth. Alexander McPherson. Journal of Crystal Growth. 110 (1991) 1-10. 

Which strategy for a protein crystallization project? C. E. Kundrot. CMLS Cellular and Molecular Life Sciences. 61 (2004) 525-536. 

Current state and prospects of macromolecular crystallography. Zbigniew Dauter. Acta Cryst. (2006). D62, 1-11. 

The impact of structural genomics: Expectations and outcomes. Steven E. Brenner et al. Science, Vol 311, Jan 20, 2006. 

Filling the niches in candidates for protein-protein interactions. Leslie Pray. May 2005, Genocis & Proteomics, 22-25. 

Life in the fast lane for protein crystallizatin and X-ray crystallography. Joseph D. Ng et al. Progress in Biophysics and Molecular Biology 88 (2005) 359-386. 

Turning protein crystallization from an art into a science. Naomi E. Chayen. Current Opinion in Structural Biology 2004, 14:577-583. 

Protein crystallization in the structural genomics era. Alexander McPherson. Journal of Structural Biology and Functional Genomics 4: 2-11, 2003. 

Insulin crystals. Chemical and biological studies on insulin crystals and insulin zinc suspensions. Jorgen Schlichtkrull. QP 951 537. Copenhagen, Ejnar Munksgaard Publishers, 1958. 

Crystallization to obtain protein-ligand complexes for structure-aided drug design. Danley. Acta Cryst. (2006). D62, 569-575. 

Strategies for structural proteomics of prokaryotes: Quantifying the advantages of studying orthologous proteins and of using both NMR and X-ray crystallography approaches. Savchenko et al. Proteins: Structure, Function, and Genetics 50:392-399 (2003). 

The prospects of protein nanocrystallography. Bodenstaff et al. Acta Cryst. (2002). D58, 1901-1906. 

The crystallization of proteins, nucleic aicds, and viruses for X-ray diffraction analysis. Alexander McPherson, Bob Cudney and Sam Patel. S. R. Fahnestock and A. Steinbuchel, eds. Biopolymers, Vol. 8, Chapter 16, 427-468. 2003. 

High-throughput crystallization and structure determination in drug discovery. Lance Stewart et al. DDT Vol. 7, No. 3 February 2002. 

Crystallogenesis of biological macromolecules. Biological, microgravity and other physicochemical aspects. R. Giege, J. Drenth, A. Ducruix, A McPherson and W. Saenger. Prog. Crystal Growth and Charact. Volume 30 pages 237-281, 1995. 

Lessons in stability from thermophile proteins. Abbas Razvi and J. Martin Scholtz. Protein Science (2006). 15:1569-1578.

Polyethylene glycol interactions with proteins. Z. Kristallogr. Suppl. 23 (2006) 613-618. 

Agarose gel facilitates enzyme crystal soaking with a ligand analog. C. Sauter, C. Balg, A. Moreno, K. Dhouib, A. Théobald-Dietrich, R. Chênevert, R. Giegé and B. Lorber. J. Appl. Cryst. (2009). 42, 279-283. 

Effect of amino acids and amino acid derivatives on crystallization of hemoglobin and ribonuclease A. Let Ito et al. J. Synchrotron Rad. (2008) 15, 316-318. 

MAD phasing using the (Ta6Br12)2+ cluster: a retrospective study. M. Jaskolski et al. Acta Cryst. (2008) D64, 595-606. 

Towards a rational approach for heavy-atom derivative screening in protein crystallography. Johnson Agniswamy, M. Gordon Joyce, Carl H. Hammer, and Peter D. Sun. Acta Cryst. (2008) D64, 354-367. 

Silver Bullets: An Alternative Strategy for Crystallizing Macromolecules. A. McPherson, B. Cudney / Journal of Structural Biology 156 (2006) 387–406 

A novel strategy for the crystallization of proteins: X-ray diffraction validation. Acta Cryst. (2007). D63, 310–318 

Using barium ions for heavy-atom derivatization and phasing of xylanase II from Trichoderma longibrachiatum. Natalia Moiseeva and 
Marc Allaire. Acta Cryst. (2007). D63, 1025–1028. 

Understanding the crystallization of an acidic protein by dilution in the ternary NaCl–2-methyl-2,4-pentanediol–H2O system. Lionel Costenaroa, Giuseppe Zaccaia, b and Christine Ebel. Journal of Crystal Growth, Volume 232, Issues 1-4, November 2001, Pages 102-113. 

Crystal engineering of the composition of pharmaceutical phases. Do pharmaceutical co-crystals represent a new path to imporved medicines? Orn Almarsson and Machael Zaworotko. Chem. Commun. 2004, 1889-1896. 

Ionic strength reducers: an efficient approach to protein purification and crystallization. Application to two Rop variants. Papanikolau et al. Acta Cryst. (2004). D60, 1334-1337. 

Effects of organic anions on the crystallization of the Ca2+-ATPase of organic sarcoplasmic reticulum. Misra et al. Biochimica et Biophysica Acta. 1077 (1991) 1-7-118. 

An overview on 2-methyl-2,4-pentanediol in crystallization and in crystals of biological macromolecules. Anand et al. Acta Cryst. (2002), D58, 1722-1728. 

Crystal structure of constitutive endothelial nitric oxide synthase: a paradigm for pterin function involving a novel metal center. Raman et al. Cell, Vol. 95, 939-950, December 23, 1998. (SDS as an additive) 

Use of glycerol, polyols and other protein structure stabilizing agents in protein crystallization. Rui Sousa. Acta Cryst. (1995). D51, 271-277. 

Use of organic cosmotropic solutes to crystallize flexible proteins: application to T7 polymerase and its complex with the inhibitor T7 lysozyme. J. Mol. Biol. (1997) 274, 748-756. 

Cocrystallizing agents for amino acids. II. The crystal structures of L-histidine-4,5-imidazoledicarboxylic acid (1:1) and L-lysine-4,5-imidazoledicarboxylic acid (1:1). Carl Henrik Gorbitz and Johanne Husdal. Acta Chemica Scandinavica 52 (1998) 218-226. 

Hexafluoroacetone hydrate as a structure modifier in proteins: characterization of a molten globule state of hen egg-white lysozyme. Surajit Bhattacharjya and Padmanabhan Balaram. Protein Science (1997), 6: 1065-1073. 

Effects of polyethylene glycol-400 at low concentrations on the long term growth of muscle phosphoglucomutase crystals from concentrated salt solutions. William J. Ray. Proteins: Structure, Function, and Genetics 14: 300-308 (1992). 

Relative effectiveness of various anions on the solubility of acidic Hypoderma lineatum collagenase at pH 7.2. Carbonnaux et al. Protein Science (1995), 4: 2123-2128. 

Ionic strength reducers: an efficient approach to protein purification and crystallization. Application to two Rop variants. Michael Kokkinidis et al. Acta Cryst. (2004). D60, 1334-1337. 

Protein stabilization by specific binding of guanidinium to a functional arginine-binding surface on an SH3 domain. Alan R. Davidson et al. Protein Science (2006) 15:162-170. 

Formation of substrate and transition-state analogue complexes in crystals of phosphoglucomutase after removing the crystallization salt. William J. Ray et al. Biochemistry. 1991 Jul 16;30(28):6875-85. 

Detection of L-lactate in polyethylene glycol solutions confirms the identity of the active-site ligand in a proline dehydrogenase structure. M. Zhang and J.J. Tanner Acta Crystallogr. (2004) D60, 985-986 .

Automatic implementation of precise grid screens: the four-corners method. Rosenberg et al, Acta Crysta. (2009), D65, 1001-1003. 

Cyclic olefin homopolymer-based microfluidics for protein crystallization and in situ X-ray diffraction. Emamzadah et al, Acta Cryst. (2009), D65, 913-920. 

Towards a ‘universal’ nucleant for protein crystallization. Emmanuel Saridakis1 and Naomi E. Chayen. Trends in Biotechnology, Volume 27, Issue 2, February 2009, Pages 99-106. 

An automated microseed matrix-screening method for protein crystallization. Allan D'Arcy, Frederic Villard and May Marsh. Acta Cryst. (2007) D63, 550-554. 

Microscale vapour diffusion for protein crystallization. J. Korczynska, T.-C. Hu, D. K. Smith, J. Jenkins, R. Lewis, T. Edwards and A. M. Brzozowski. Acta Cryst. (2007). D63, 1009-1015. 

A method for the general identification of protein crystals in crystallization experiments using a noncovalent fluorescent dye. Groves MR, Muller IB, Kreplin X, Muller-Dieckmann J. Acta Crystallogr D Biol Crystallogr. 2007 Apr;63(Pt 4):526-35. 

Trace fluorescent labeling for high-throughput crystallography. Forsythe E, Achari A, Pusey ML. Acta Crystallogr D Biol Crystallogr. 2006 Mar;62(Pt 3):339-46. 

A complete microfluidic screening platform for rational protein crystallization. Hansen et al. J. Am. Chem. Soc. 2007, 129, 454-455. 

Nanoliter microfluidic hybrid method for simultaneous screening and optimization validated with crystallization of membrane proteins. Ismagilov et al. PNAS December 19, 2006 Volume 103 No 51 19243-19248. 

An apparatus for electric-field-induced protein crystallization. M. I. Al-Haq, E. Lebrasseur, W.-K. Choi, H. Tsuchiya, T. Torii, H. Yamazaki and E. Shinohara. J. Appl. Cryst. (2007). 40, 199-201 

The open-access high-throughput crystallization facility at EMBL Hamburg. J. Mueller-Dieckmann. Acta Cryst. (2006). D62, 1446-1452. 

A serendipitous discovery that in situ proteolysis is essential for the crystallization of yeast CPSF-100 (Ydh1p). Corey R. Mandel, Damara Gebauer, Hailong Zhang and Liang Tong. Acta Cryst. (2006). F62, 1041–1045 

Automated streak-seeding with micromachined silicon tools. A. Georgiev, S. Vorobiev, W. Edstrom, T. Song, A. Laine, J. Hunt and P. Allen. Acta Cryst. (2006). D62, 1039-1045 

SPINE high-throughput crystallization, crystal imaging and recognition techniques: current state, performance analysis, new technologies and future aspects. Ian M. Berry, O. Dym, R.M. Ensouf, K. Harlos, R. Meged, A. Perrakis, J.L. Sussman, T.S. Walter, J. Wilson, and Albrecht Messerschmidt. Acta Cryst. (2006). D62, 1137-1149. 

Heterogeneous and epitaxial nucleation of protein crystals on mineral surfaces. Alexander McPherson and Paul Shlichta. Science, 22 January 1988, Volume 239, pp.385-387. 

Picolitra-scale crystallization of membrane proteins. Vadim Cherezov and Martin Caffrey. J. Appl. Cryst. (2006). 39, 604-606. 

Use of plastic capillaries for macromolecular crystallization. Potter et al. J. Appl. Cryst. (2004). 37, 500-501. 

Structural study of the type-II 3-dehydroquinate dehydratase from Actinobacillus pleuropneumooniae. Maes et al. Acta Cryst. (2004). D60, 463-471. (gel accupuncture) 

Microseed matrix screening to improve crystals of yeast cytosine deaminase. Ireton and Stoddard. Acta Cryst. (2004). D60, 601-605. 

Protein nano-crystallogenesis. Kuil et al. Enzyme and Microbial Technology 30 (2002) 262-265. 

Automatic classificaiton of protein crystallization images using a curve-tracking algorithm. Bern et al. J. Appl. Cryst. (2004). 37, 279-287. 

A novel approach to crystallizing proteins with temperature induction method: GrpE protein from Thermus thermophilus. Kitano et al. Journal of Crystal Growth 186 (1998) 456-460. 

Accelerate protein crystal growth by protein thin film template. Pechkova and Nicolini. Journal of Crystal Growth 231 (2001) 599-601. 

Development of a technology for automation and miniaturization of protein crystallization. Mueller et al. Journal of Biotechnology 85 (2001) 7-14. 

Practical aspects of using the microbatch method in screening conditions for protein crystallization. D'Arcy et al. Methods 34 (2004) 323-328. 

Porous silicon: an effective nucleation-indusing material for protein crystallization. Chayen et al. J. Mol. Biol. (2001) 312, 591-595. 

Interface sliding as illustrated by the multiple quaternary structures of liganded hemoglobin. Mueser et al. Biochemist.

New paradigm for macromolecular crystallography experiments at SSRL: automated crystal screening and remote data collection. S. Michael Soltis et al. Acta Cryst. (2008) D64, 1210-1221 

Hyperquenching for protein crystallography. Matthew Warkentin, Viatcheslav Berejnov, Naji Husseini, Robert Thorne. J. Appl. Cryst. (2006). 39, 805-811 

Radiation damage in macromolecular cryocrystallography. Ravelli, Raimond Bg; Garman, Elspeth F. Curr Opin Struct Biol Volume 16, Number 5, 2006 Oct, PP 624-9. 

Quantifying X-ray radiation damage in protein crystals at cryogenic temperatures. J. Kmetko, N. S. Husseini, M. Naides, Y. Kalinin and R. E. Thorne. Acta Cryst. (2006). D62, 1030-1038 

An automated temperature-monitoring system for dry-shippers. F. Ridoutt, C. Mueller-Dieckmann, P. A. Tucker and M. S. Weiss. J. Appl. 
Cryst., 2004, 37,477-480. 

Effects of cryoprotectant concentration and cooling rate on vitrification of aqueous solutions. Robert E. Thorne et al. J. Appl. Cryst. (2006) 39, 244-251. 

Cryocooling and radiation damage in macromolecular crystallography. Elspeth Garman et al. Acta Cryst. (2006). D62, 32-47. 

A possible additional role of mineral oil in successful flash cooling. John. M. Wiencek et al. J. Appl. Cryst. (2005). 38, 945-950. 

A novel cryoprotection scheme for enhancing the diffraction of crystals of recombinant cytochrome ba3 oxidase from Thermus thermophilus. David Stout et al. Acta Cryst. (2005). D61, 340-343. 

Cool crystals: macromolecular cryocrystallography and radiation damage. Elspeth Garman. Current Opinion in Structural Biology 2003, 13:545-551. 

Temperature characteristics of crystal storage devices in a CP100 dry shipping dewar. Robin Leslie Owen, Mike Pritchard and Elspeth Garman. J. Appl. Cryst. (2004). 37, 1000-1003. 

Sample size: an important parameter in flash-cooling macromolecular crystallization solutions. C. Schall et al. J. Appl. Cryst. (2005). 38, 412-419. 

Cool data: quantity and quality. Elspeth Garman. Acta Cryst. (1999). D55, 1641-1653. 

Cryocrystallography with oil - an old idea revived. Riboldi-Tunnicliffe and Hilgenfeld. J. Appl. Cryst. (1999). 32, 1003-1005. 

A cryocooling technique for protein crystals grown by dialysis from volatile solvents. Fernandez et al. J. Appl. Cryst. (2000). 33, 168-171. 

J. Appl. Cryst. (1996). 29, 584-587 [ doi:10.1107/S0021889896004190 ] 
Glycerol concentrations required for cryoprotection of 50 typical protein crystallization solutions E. F. Garman and E. P. Mitchell 

Acta Cryst. (2008). D64, 287-301 [ doi:10.1107/S0907444907067613 ] 
Glycerol concentrations required for the successful vitrification of cocktail conditions in a high-throughput crystallization screen R. Kempkes, E. Stofko, K. Lam and E. H. Snell 

MyCrystals - a simple visual data management program for laboratory-scale crystallization experiments. M. N. Løvgreen, M. Løvgreen, H. E. M. Christensen and P. Harris. J. Appl. Cryst. (2009). 42. Synopsis: MyCrystals is a simple user-friendly program developed to display and sort crystal images on the basis of the entered crystallization conditions. 

MPCD: a new interactive on-line crystallization data bank for screening strategies. M. Charles, S. Veesler and F. Bonneté. Acta Cryst. (2006). D62, 1311-1318.

Selecting optimum eukaryotic integral membrane proteins for structure determination by rapid expression and solubilization screening. Li M, Hays FA, Roe-Zurz Z, Vuong L, Kelly L, Ho CM, Robbins RM, Pieper U, O'Connell JD 3rd, Miercke LJ, Giacomini KM, Sali A, Stroud RM. J Mol Biol. 2009 Jan 23;385(3):820-30. 

A strategy for identification and quantification of detergents frequently used in the purification of membrane proteins. Laura R. Eriks, June A. 
Mayor and Ronald S. Kaplan. Analytical Biochemistry Volume 323, Issue 2, 15 December 2003, Pages 234-241. 

A high-throughput method for membrane protein solubility screening: The ultracentrifugation dispersity sedimentation assay 
Daniel A.P. Gutmann, Eiichi Mizohata, Simon Newstead, Sebastian Ferrandon, Peter J.F. Henderson, Hendrik W. van Veen, and Bernadette Byrne. Protein Sci 2007 16: 1422-1428 

When worlds colloid. Michael C. Weiner. Protein Science (2006), 15:2679-2681. 

Effects of additives on surfactant phase behavior relevant to bacteriorhodopsin crystallization. Bryan Berger, Colleen Gendron, Abraham Lenhoff and Eric Kaler. Protein Science (2006), 15:2682-2696. 

A defined protein-detergent-lipid complex for crystallization of integral membrane 
proteins: The cytochrome b6f complex of oxygenic photosynthesis. Zhang H, Kurisu G, Smith JL, Cramer WA. Proc Natl Acad Sci U S A. 2003 Apr 29;100(9):5160-3.

Schultz, S. C., Shields, G. C. & Steitz, T. A. (1990). Crystallization of Escherichia coli catabolite gene activator protein with its DNA binding site. J. Mol. Biol. 213, 159–166. 

Tan et al, Crystallization of the Yeast MATa2/MCM1/DNA Ternary Complex: General Methods and Principles for Protein/DNA Cocrystallization J. Mol. Biol. (2000) 297, 947-959. 

Cannone et al, Crystallization of bFGF-DNA aptamer complexes using a 

Sparse Matrix designed for protein–nucleic acid complexes Journal of Crystal Growth, 2001 232 (2001) 409–417. 

Structure of a compact conformation of linear diubiquitin. Ahmed Rohaim, Masato Kawasaki,Ryuichi Kato, Ivan Dikicd and Soichi Wakatsuki. Acta Cryst. (2012). D68, 102–108. Hampton Research Crystal Screen reagent 6 Crystal Screen condition No. 6 consisting of 0.2 M magnesium chloride hexahydrate, 0.1 M Tris–HCl pH 8.5, 30%(w/v) polyethylene glycol 4000 produced preliminary crystals. Crystals optimized using 30% PEG 4000, 0.1 M Tris–HCl, 0.1 M MgCl2, 30% dioxane, 30% 2-propanol at pH 7.5. Crystals diffracted to 2.2 angstrom. 

Structure of Escherichia coli BamD and its functional implications in outer membrane protein assembly. Cheng Dong, Hai-Feng Hou, Xue Yang, Yue-Quan Shenb and Yu-Hui Dong. Acta Cryst. (2012). D68, 95–101 doi:10.1107. Hampton Research PEG/Ion screen utilized to produce preliminary crystals (20%(w/v) PEG 3350 and 0.20 M potassium iodide). Reagent optimized to 27%(w/v) PEG 3350 and 0.28 M potassium iodide. Structure solved to 2.6 angstrom. 

Structural insights into human Kif7, a kinesin involved in Hedgehog signalling. Klejnot & Kozielski. Acta Cryst. (2012). D68, 154–159. Hampton Research PEG/Ion screen reagent containing 0.2 M lithium acetate, 20%(w/v) PEG 3350 produces preliminary crystals.

Lysozyme contamination facilitates crystallization of a heterotrimeric cortactin-Arg-lysozyme complex. W. Liu, S. M. MacGrath, A. J. Koleske and T. J. Boggon. Acta Cryst. (2012). F68, 154-158 [ doi:10.1107/S1744309111056132 ]. Synopsis: An unusual case of trace amounts of a contaminating protein facilitating formation of a heterotrimeric protein complex. 

Genetic Construct Design and Recombinant Protein Expression for Structural Biology. Suzanne C. Edavettal, Michael J. Hunter and Ronald V. Swanson. Structure-Based Drug Discovery Methods In Molecular Biology™, 2012, Volume 841, 29-47, DOI: 10.1007/978-1-61779-520-6_2. 

High-throughput protein crystallization on the World Community Grid and the GPU. Yulia Kotseruba, Christian A Cumbaa and Igor Jurisica. J. Phys.: Conf. Ser. 341 012027 doi:10.1088/1742-6596/341/1/012027. 

X-ray Diffraction of Protein Crystal Grown in a Nano-liter Scale Droplet in a Microchannel and Evaluation of Its Applicability. Maeki et al. Analytical Sciences Vol. 28 (2012) , No. 1 p.65. 

CRYSpred: Accurate Sequence-Based Protein Crystallization Propensity Prediction Using Sequence-Derived Structural Characteristics. MarcinJ. Mizianty; A. Kurgan, Lukasz. Protein and Peptide Letters, Volume 19, Number 1, January 2012 , pp. 40-49(10). 

Enhancement of crystallization with nucleotide ligands identified by dye-ligand affinity chromatography. Heungbok Kim, Cecelia Webster, Justin K. M. Roberts, Juthamas Kositsawat, Li-Wei Hung, Thomas C. Terwilliger and Chang-Yub Kim. Journal of Structural and Functional Genomics DOI: 10.1007/s10969-012-9124-8. 

Lipidic phase membrane protein serial femtosecond crystallography. Johannson et al. Nature Methods(2012)doi:10.1038/nmeth.1867. 

On the need for an international effort to capture, share and use crystallization screening data. J. Newman, E. E. Bolton, J. Mueller-Dieckmann, V. J. Fazio, D. T. Gallagher, D. Lovell, J. R. Luft, T. S. Peat, D. Ratcliffe, R. A. Sayle, E. H. Snell, K. Taylor, P. Vallotton, S. Velanker and F. von Delft. Acta Cryst. (2012). F68, 253-258 [doi:10.1107/S1744309112002618] 

Improvement in Quality of Protein Crystals Grown in a High Magnetic Field Gradient. Akira Nakamura, Jun Ohtsuka, Ken-ichi Miyazono, Akihiro Yamamura, Keiko Kubota, Ryoichi Hirose, Noriyuki Hirota, Mitsuo Ataka, Yoriko Sawano, and Masaru Tanokura. Crystal Growth & Design. Publication Date (Web): 27 Feb 2012 (Article) DOI: 10.1021/cg2007414. 

Control of Gibbs free energy relationship between hen egg white lysozyme polymorphs under application of an external alternating current electric field. Y. Tomita, H. Koizumi, S. Uda, K. Fujiwara and J. Nozawa. J. Appl. Cryst. (2012). 45, 207-212 [ doi:10.1107/S002188981200249X ] 

High-throughput biological small-angle X-ray scattering with a robotically loaded capillary cell. S. S. Nielsen, M. Møller and R. E. Gillilan. J. Appl. Cryst. (2012). 45, 213-223 [doi:10.1107/S0021889812000957 ] 

A fibre-based crystal mounting technique for protein Cryocrystallography. M. Sugahara. J. Appl. Cryst. (2012). 45, 362-366 [ doi:10.1107/S002188981200756X ] 

Computational design of a protein crystal. Christopher J. Lancia, Christopher M. MacDermaida, Seung-gu Kanga, Rudresh Acharyab, Benjamin Northb, Xi Yanga, X. Jade Qiub, William F. DeGrado, and Jeffery G. Savena. Published online before print April 25, 2012, doi: 10.1073/pnas.1112595109 PNAS April 25, 2012. 

Charge-controlled metastable liquid–liquid phase separation in proteinsolutions as a universal pathway towards crystallization. Fajun Zhang, Roland Roth, Marcell Wolf, Felix Roosen-Runge, Maximilian W. A. Skoda, Robert M. J. Jacobs , Michael Stzuckiand Frank Schreiber. Soft Matter, 2012, 8, 1313-1316. DOI: 10.1039/C2SM07008A 
Summary: We demonstrate that a metastable liquid–liquid phase separation (LLPS) in protein aqueous solutions can be induced by multivalent metal ions at room temperature. We determine the salt andprotein partitioning in the two coexisting phases. The structure factor obtained by small angle X-ray scattering provides direct evidence for a short-ranged attraction, which leads to the metastability of the LLPS. An extended phase diagram with three control parameters (temperature, protein and salt concentration) provides a conclusive physical picture consistent with a criterion for the second virial coefficient. The presented isothermal control mechanism of the phase behavior opens new perspectives for the understanding of controlled phase behavior in nature. Furthermore, we discuss the application of this framework in predicting and optimizing conditions for protein crystallization. 

In vivo protein crystallization opens new routes in structural biology. Rudolf Koopmann, Karolina Cupelli, Lars Redecke, Karol Nass, Daniel P DePonte, Thomas A White, Francesco Stellato, Dirk Rehders, Mengning Liang, Jakob Andreasson, Andrew Aquila, Sasa Bajt, Miriam Barthelmess, Anton Barty, Michael J Bogan, Christoph Bostedt, Sébastien Boutet, John D Bozek, Carl Caleman, Nicola Coppola, Jan Davidsson, R Bruce Doak, Tomas Ekeberg, Sascha W Epp, Benjamin Erk et al. Nature Methods 9, 259–262 (2012) doi:10.1038/nmeth.1859. Summary: Protein crystallization in cells has been observed several times in nature. However, owing to their small size these crystals have not yet been used for X-ray crystallographic analysis. We prepared nano-sized in vivo–grown crystals of Trypanosoma brucei enzymes and applied the emerging method of free-electron laser-based serial femtosecond crystallography to record interpretable diffraction data. This combined approach will open new opportunities in structural systems biology. 

Predicting Protein Crystallizability and Nucleation. Sánchez-Puig N, Sauter C, Lorber B, Giegé R, Moreno A. Protein Pept Lett. 2012 Apr 4. Summary: The outcome of protein crystallization attempts is often uncertain due to inherent features of the protein or to the crystallization process that are not fully under control of the experimentalist. The aim of this contribution is to propose user-friendly tools that can increase the success rate of a protein crytallization project. Different bioinformatic approaches to predict the crystallization feasibility (before any crystallization attempts are undertaken) and a novel approach to assess the nucleation process of a given protein is proposed. Practical examples illustrate these two points. 

Growth Rates of Protein Crystals. Jeremy D. Schmit and Ken Dill. J. Am. Chem. Soc., 2012, 134 (9), pp 3934–3937 DOI: 10.1021/ja207336r. Summary: Protein crystallization is important for structural biology. The rate at which a protein crystallizes is often the bottleneck in determining the protein’s structure. Here, we give a physical model for the growth rates of protein crystals. Most materials crystallize faster under stronger growth conditions; however, protein crystallization slows down under the strongest conditions. Proteins require a crystallization slot of ‘just right’ conditions. Our model provides an explanation. Unlike simpler materials, proteins are orientationally asymmetrical. Under strong conditions, protein molecules attempt to crystallize too quickly, in wrong orientations, blocking surface sites for more productive crystal growth. The model explains the observation that increasing the net charge on a protein increases the crystal growth rate. The model predictions are in good agreement with experiments on the growth rates of tetragonal lysozyme crystals as a function of pH, salt concentration, temperature, and protein concentration. 
High-throughput protein crystallization on the World Community Grid and the GPU. Yulia Kotseruba, Christian A Cumbaa and Igor Jurisica. Journal of Physics: Conference Series Volume 341 conference Yulia Kotseruba et al 2012 J. Phys.: Conf. Ser. 341 012027 doi:10.1088/1742-6596/341/1/012027. Summary: We have developed CPU and GPU versions of an automated image analysis and classification system for protein crystallization trial images from the Hauptman Woodward Institute's High-Throughput Screening lab. The analysis step computes 12,375 numerical features per image. Using these features, we have trained a classifier that distinguishes 11 different crystallization outcomes, recognizing 80% of all crystals, 94% of clear drops, 94% of precipitates. The computing requirements for this analysis system are large. The complete HWI archive of 120 million images is being processed by the donated CPU cycles on World Community Grid, with a GPU phase launching in early 2012. The main computational burden of the analysis is the measure of textural (GLCM) features within the image at multiple neighbourhoods, distances, and at multiple greyscale intensity resolutions. CPU runtime averages 4,092 seconds (single threaded) on an Intel Xeon, but only 65 seconds on an NVIDIA Tesla C2050. We report on the process of adapting the C++ code to OpenCL, optimized for multiple platforms. 

Randomness in Crystallization of Proteins from Staphylococcus Aureus. Yan S, Wu G. Protein Pept Lett. 2012 Apr 4. Summary: Of many factors affecting protein crystallization, randomness in proteins has been given less attention although highly structured proteins would be at low entropy state. The factors, which impact on protein crystallization, are almost exclusively related to non-random amino acid properties such as physiochemical properties of amino acids. In this study, we used logistic regression and neural network to model the success rate of crystallization of 420 proteins from Staphylococcus aureus with each of non-random and random amino acid properties in order to determine whether randomness in a protein plays a role in the crystallization process. The results show that randomness is indeed involved in the crystallization process, and this rationale would enrich our knowledge on crystallization process and enhance our ability to crystallize more important proteins. 

The structure of an orthorhombic crystal form of a `forced reduced' thiol peroxidase reveals lattice formation aided by the presence of the affinity tag. K. S. H. Beckham, O. Byron, A. J. Roe and M. Gabrielsen. Acta Cryst. (2012). F68, 522-526 [ doi:10.1107/S1744309112011487 ]. Summary: The crystal structure of the C61S mutant of Tpx from E. coli is reported at 1.97 AA resolution. A brief structural comparison with homologues is presented. 

Use of differential scanning fluorimetry to optimize the purification and crystallization of PLP-dependent enzymes. T. W. Geders, K. Gustafson and B. C. Finzel. Acta Cryst. (2012). F68, 596-600 [ doi:10.1107/S1744309112012912 ]. Summary: Differential scanning fluorimetry (DSF) is a practical and accessible technique that allows the assessment of multiphasic unfolding behavior resulting from subsaturating binding of ligands. Multiphasic unfolding is indicative of a heterogenous protein solution, which frequently interferes with crystallization and complicates functional characterization of proteins of interest. Along with UV-Vis spectroscopy, DSF was used to guide purification and crystallization improvements for the pyridoxal 5'-phosphate (PLP) dependent transaminase BioA from Mycobacterium tuberculosis. The incompatibility of the primary amine-containing buffer 2-amino-2-(hydroxymethyl)-1,3-propanediol (Tris) and PLP was identified as a significant contributor to heterogeneity. It is likely that the utility of DSF for ligand-binding assessment is not limited to the cofactor PLP but will be applicable to a variety of ligand-dependent enzymes. 

On-column ligand exchange for structure-based drug design: a case study with human 11[beta]-hydroxysteroid dehydrogenase type 1. W. Qin, R. A. Judge, K. L. Longenecker, L. R. Solomon and J. E. Harlan. Acta Cryst. (2012). F68, 601-605 
[ doi:10.1107/S1744309112010172 ]. Summary: An on-column ligand- and detergent-exchange method was developed to obtain ligand-protein complexes for an adamantane series of compounds with 11[beta]-HSD1 after a variety of other complexation methods had failed. An interesting byproduct of the method was the observation of artificial trimers in the crystal structures. 

Analysis of polycrystallinity in hen egg-white lysozyme using a pnCCD. S. Send, A. Abboud, W. Leitenberger, M. S. Weiss, R. Hartmann, L. Strueder and U. Pietsch. J. Appl. Cryst. (2012). 45, 517-522 [ doi:10.1107/S0021889812015038 ] Synopsis: An analysis of polycrystallinity in tetragonal hen egg-white lysozyme by means of energy-dispersive Laue diffraction techniques using a pnCCD detector is presented. 

Protonation-state determination in proteins using high-resolution X-ray crystallography: effects of resolution and completeness. S. J. Fisher, M. P. Blakeley, M. Cianci, S. McSweeney and J. R. Helliwell. Acta Cryst. (2012). D68, 800-809 [ doi:10.1107/S0907444912012589 ]. Synopsis: A bond-distance analysis to determine the protonation states of ionizable amino acids has been made for trypsin at 1.2 Å resolution, subtilisin at 1.26 Å resolution and lysozyme at 0.65 Å resolution. This was effective for Asp and Glu but not for His. 

Outrunning free radicals in room-temperature macromolecular crystallography. R. L. Owen, D. Axford, J. E. Nettleship, R. J. Owens, J. I. Robinson, A. W. Morgan, A. S. Doré, G. Lebon, C. G. Tate, E. E. Fry, J. Ren, D. I. Stuart and G. Evans. Acta Cryst. (2012). D68, 810-818 [ doi:10.1107/S0907444912012553 ]. Synopsis: A systematic increase in lifetime is observed in room-temperature protein and virus crystals through the use of reduced exposure times and a fast detector. 

Crystallization and preliminary X-ray diffraction studies of the GhKCH2 motor domain: alteration of pH significantly improved the quality of the crystals. X. Qin, Z. Chen, T. Xu, P. Li and G. Liu. Acta Cryst. (2012). F68, 798-801 [ doi:10.1107/S1744309112016351 ]. Synopsis: The GhKCH2 motor domain was crystallized and the pH of the crystallization buffer was shown to have a significant effect on the crystal morphology and diffraction quality. 

Crystallization of SHARPIN using an automated two-dimensional grid screen for optimization. B. Stieglitz, K. Rittinger and L. F. Haire. Acta Cryst. (2012). F68, 816-819 [ doi:10.1107/S1744309112022208 ]. Synopsis: The expression, purification and crystallization of an N-terminal fragment of SHARPIN are reported. Diffraction-quality crystals were obtained using a two-dimensional grid-screen seeding technique. 


Combining in situ proteolysis and mass spectrometry to crystallize Escherichia coli PgaB. D. J. Little, J. C. Whitney, H. Robinson, P. Yip, M. Nitz and P. L. Howell. Acta Cryst. (2012). F68, 842-845 doi:10.1107/S1744309112022075. Synopsis: Construct engineering and crystallization of E. coli PgaB using in situ proteolysis and mass spectrometry is reported. 


Rehydratable gel for rapid loading of nanoliter solution and its application in protein crystallization. Yuefang Li , Dameng Guo and Bo Zheng. RSC Adv., 2012, 2, 4857-4863 DOI: 10.1039/C2RA20511D. Synopsis: In the work presented, rehydratable polyacrylamide gel is introduced as a medium to uptake and store nanoliter protein solutions in microwells for multiplex bioanalysis. The polyacrylamide gel, produced and stored in the microwells, shrank by 97% upon dehydration and could be reversibly rehydrated to 95% of the initial volume by absorbing aqueous solution. We employed the rehydratable gel to load aqueous solutions of different proteins with molecular weights in the range of 14.7–250 kDa. The protein loading occurred simultaneously with the gel rehydration and reached saturated state in 5 min. The relative protein concentrations in the gel ranged from 92% to 53%, depending on the molecular weight of the proteins. Particularly, the rehydratable gel had a much higher protein loading efficiency than the fresh gel. We applied the protein-carrying gel to the crystallization of four model proteins in the microwells and produced diffraction-quality protein crystals. The rehydratable gel is simple to fabricate, efficient to load with protein, and has good capacity for storing the protein solutions in microwells with minimal dilution effects on the protein solution. The rehydratable gel incorporated microwell chip should be useful in multiplex analysis that requires small sample consumption and high throughput. 


Charge-controlled metastable liquid–liquid phase separation in protein solutions as a universal pathway towards crystallization. Fajun Zhang , Roland Roth , Marcell Wolf , Felix Roosen-Runge , Maximilian W. A. Skoda , Robert M. J. Jacobs , Michael Stzucki and Frank Schreiber. Soft Matter, 2012, 8, 1313-1316, DOI: 10.1039/C2SM07008A. Synopsis: We demonstrate that a metastable liquid–liquid phase separation (LLPS) in protein aqueous solutions can be induced by multivalent metal ions at room temperature. We determine the salt and protein partitioning in the two coexisting phases. The structure factor obtained by small angle X-ray scattering provides direct evidence for a short-ranged attraction, which leads to the metastability of the LLPS. An extended phase diagram with three control parameters (temperature, protein and salt concentration) provides a conclusive physical picture consistent with a criterion for the second virial coefficient. The presented isothermal control mechanism of the phase behavior opens new perspectives for the understanding of controlled phase behavior in nature. Furthermore, we discuss the application of this framework in predicting and optimizing conditions for protein crystallization. 


Hofmeister effects of ionic liquids in protein crystallization: Direct and water-mediated interactions. Magdalena Kowacz , Abhik Mukhopadhyay , Ana Luísa Carvalho , José M. S. S. Esperança , Maria J. Romão and Luís Paulo N. Rebelo. CrystEngComm, 2012, Advance Article DOI: 10.1039/C2CE25129A. Synopsis: We have performed experiments on the crystallization of two low molecular weight, positively charged proteins, lysozyme and ribonuclease A, using ionic liquids as either crystallization additives or, in particular cases, as precipitating agents. The ionic liquids (ILs) have been ordered according to their salting-in/out ability and the relative position of these ionic liquids in this ranking has been rationalized by considering their hydration properties (positive–negative, hydrophobic–hydrophilic). The ability to screen the effective charge of cationic proteins and aid protein nucleation (salting-out) has been shown to be superior for large polarizable anions with low charge density, negatively hydrated-Cl-, Br-, [SCN]-, methane-[C1SO3]- and ethanesulfonates [C2SO3]-, than for anions with a relatively stable hydration shell, positively hydrated-lactate [Lac]-, butylsulfonate [C4SO3]- and acetate [Ac]-. Upon increasing the background salt concentration, where electrostatic interactions are already effectively screened, the ability of the IL ions to stabilize proteins in solution (salting-in) has been shown to increase as the ions are likely to migrate to the non-polar protein surface and lower protein–water interfacial tension. This tendency is enhanced as the focus moves from those ions with positively hydrated hydrophilic compartments (e.g. [Ac]-) to those with negatively hydrated groups (e.g. [C1SO3]-) and the prevailing hydrophobic hydration (e.g. [C4SO3]-). The observed inversion in the relative effect of ILs on protein crystallization with increasing ionic strength of the aqueous media has been interpreted as the differing effects of ion adsorption: charge screening and interfacial tension modification. Moreover, this work can further help in our understanding of the influence of ionic liquids on conformational changes of biomacromolecules in solution. Identification of the specific incorporation sites for choline and acetate ions, localized in two lysozyme crystals grown in pure IL solutions without any buffer or inorganic precipitant, can give us some insight into the role of the ionic liquid ions in protein structure development. 
Nanolitre-scale crystallization using acoustic liquid-transfer technology. A. G. Villasenor, A. Wong, A. Shao, A. Garg, T. J. Donohue, A. Kuglstatter and S. F. Harris. Acta Cryst. (2012). D68, 893-900 doi:10.1107/S0907444912016617. Synopsis: Acoustic droplet ejection achieves precise, tipless, non-invasive transfer of diverse aqueous solutions, enabling nanolitre-scale crystallization trials. The rapid and scalable technique demonstrated successful crystal growth with diverse targets in drop volumes as small as 20 nl. 


A universal indicator dye pH assay for crystallization solutions and other high-throughput applications. J. Newman, R. A. Sayle and V. J. Fazio. Acta Cryst. (2012). D68, 1003-1009 doi:10.1107/S0907444912018768. Synopsis: A rapid plate-based pH assay has been developed that takes advantage of the automation available in a protein-crystallization laboratory. 

Proline: Mother Nature's cryoprotectant applied to protein crystallography. T. A. Pemberton, B. R. Still, E. M. Christensen, H. Singh, D. Srivastava and J. J. Tanner. Acta Cryst. (2012). D68, 1010-1018 doi:10.1107/S0907444912019580. Synopsis: The amino acid L-proline is shown to be a good cryoprotectant for protein crystals. Four examples are provided; the range of proline used for cryoprotection is 2.0-3.0 M. 


Reduction of radiation damage and other benefits of short wavelengths for macromolecular crystallography data collection. R. Fourme, V. Honkimaeki, E. Girard, K. Medjoubi, A.-C. Dhaussy 
and R. Kahn. J. Appl. Cryst. (2012). 45, 652-661 doi:10.1107/S0021889812019164. Synopsis: X-ray photons with energy higher than usual improve both the number and the quality of diffraction data from a given macromolecular crystal. 


An anti-settling sample delivery instrument for serial femtosecond Crystallography. L. Lomb, J. Steinbrener, S. Bari, D. Beisel, D. Berndt, C. Kieser, M. Lukat, N. Neef and R. L. Shoeman. J. Appl. Cryst. (2012). 45, 674-678 doi:10.1107/S0021889812024557. Synopsis: A simple and robust instrument, which overcomes the crystal settling that impairs serial femtosecond crystallography experiments, is described. 



A gradual desiccation method for improving the efficiency of protein crystallization screening. Q.-Q. Lu, X.-Z. Xie, R.-Q. Chen, Z.-Q. Wu, Q.-D. Cheng, P. Shang and D.-C. Yin. J. Appl. Cryst. (2012). 45, 758-765 doi:10.1107/S0021889812025757. Synopsis: A modification to the vapor diffusion protein crystallization method, named the gradual desiccation method, is reported. It was found that this method can significantly enhance crystallization of proteins. 


Fine-needle capillary mounting for protein microcrystals. M. Makino, I. Wada, N. Mizuno, K. Hirata, N. Shimizu, T. Hikima, M. Yamamoto and T. Kumasaka. J. Appl. Cryst. (2012). 45, 785-788 doi:10.1107/S0021889812024545. Synopsis: It is demonstrated that a cryocrystallographic mounting method using a fine-needle capillary is suitable for protein microcrystals. 

Crystallization of Pseudomonas aeruginosa AmrZ protein: development of a comprehensive method for obtaining and optimization of protein-DNA crystals. E. E. Pryor, D. J. Wozniak and T. Hollis. Acta Cryst. (2012). F68, 985-993 doi:10.1107/S1744309112025316. Synopsis: Crystallization of the complex of the transcription factor AmrZ with DNA was accomplished through the combination of established and newly developed methods. Here, a general method to obtain and optimize crystals of protein-DNA complexes consisting of these combined procedures is described. 

Single-drop optimization of protein crystallization. A. Meyer, K. Dierks, D. Hilterhaus, T. Klupsch, P. Mühlig, J. Kleesiek, R. Schöpflin, H. Einspahr, R. Hilgenfeld and C. Betzel. Acta Cryst. (2012). F68, 994-998 doi:10.1107/S1744309112024074. Synopsis: A device has been developed to optimize crystal-growth conditions by experiments in a single drop. 

Contamination from an affinity column: an encounter with a new villain in the world of membrane-protein crystallization. P. Panwar, A. Deniaud and E. Pebay-Peyroula. Acta Cryst. (2012). D68, 1272-1277 [ doi:10.1107/S090744491202639X ]. Synopsis: Small crystals of contaminant protein led to the structure at 1.9 Å resolution of Strep-Tactin in complex with desthiobiotin. Trace amounts of Strep-Tactin were observed to be eluted from a Strep-Tactin column using several detergents, illustrating their possible role in the contamination when crystallizing membrane proteins. 

Crystallization, dehydration and experimental phasing of WbdD, a bifunctional kinase and methyltransferase from Escherichia coli O9a. G. Hagelueken, H. Huang, K. Harlos, B. Clarke, C. Whitfield and J. H. Naismith. Acta Cryst. (2012). D68, 1371-1379 [ doi:10.1107/S0907444912029599 ]. Synopsis: The optimization of WbdD crystals using a novel dehydration protocol and experimental phasing at 3.5 Å resolution by cross-crystal averaging followed by molecular replacement of electron density into a non-isomorphous 3.0 Å resolution native data set are reported. 

CrystalDirect: a new method for automated crystal harvesting based on laser-induced photoablation of thin films. F. Cipriani, M. Röwer, C. Landret, U. Zander, F. Felisaz and J. A. Márquez. Acta Cryst. (2012). D68, 1393-1399 [ doi:10.1107/S0907444912031459 ]. Synopsis: A new crystallization support and associated method designed to fully automate the process of crystal harvesting is presented. In this system, crystals are grown on an ultrathin film that can be excised by laser photoablation and directly attached to a pin for X-ray diffraction experiments. 

High-throughput counter-diffusion capillary crystallization and in situ diffraction using high-pressure freezing in protein crystallography. M. Kurz, B. Blattmann, A. Kaech, C. Briand, P. Reardon, U. Ziegler and M. G. Gruetter. J. Appl. Cryst. (2012). 45, 999-1008 [ doi:10.1107/S0021889812034061 ]. Synopsis: Counter-diffusion crystallization using a new capillary crystallization format (CrystalHarp) is described. It allows low-temperature in situ diffraction data collection and is applicable for the novel high-pressure vitrification method, which allows data collection without the addition of cryoprotectant. 

Digital topography with an X-ray CCD camera for characterizing perfection in protein crystals. K. Wako, K. Kimura, Y. Yamamoto, T. Sawaura, M. Shen, M. Tachibana and K. Kojima. J. Appl. Cryst. (2012). 45, 1009-1014 [ doi:10.1107/S0021889812032049 ]. Synopsis: The defects in tetragonal hen egg-white lysozyme crystals were characterized by digital X-ray topography using an X-ray CCD camera and conventional X-ray topography using X-ray film. 

Non-robotic high-throughput setup for manual assembly of nanolitre vapour-diffusion protein crystallization screens. R. Skrabana, O. Cehlar and M. Novak. J. Appl. Cryst. (2012). 45, 1061-1065 [ doi:10.1107/S0021889812036527 ]. Synopsis: A method for reproducible manual formation of nanolitre protein crystallization drops in high-throughput format is described. The method saves precious protein material without sacrificing the effectiveness of the screening process, can produce diffraction-quality crystals and is easily adoptable by ordinary crystallography laboratories. 

Enhancing the volume and the optical quality of hen egg-white lysozyme crystals by coupling the salt concentration gradient crystallization method with a magnetic field. E. Magay, S. J. Cho and S. A. Kim. J. Appl. Cryst. (2012). 45, 1066-1068 [ doi:10.1107/S0021889812036060 ]. Synopsis: The effect of coupling the salt concentration gradient crystallization method with the use of the paramagnetic salt MnCl2 and a magnetic field is reported. The use of a simple magnetic device is shown to have a significant effect on hen egg-white lysozyme crystal growth. 

Improved crystallization of Escherichia coli ATP synthase catalytic complex (F1) by introducing a phosphomimetic mutation in subunit. A. Roy, M. L. Hutcheon, T. M. Duncan and G. Cingolani. Acta Cryst. (2012). F68, 1229-1233 [ doi:10.1107/S1744309112036718 ]. Synopsis: A phosphomimetic mutation in subunit dramatically increases reproducibility for crystallization of Escherichia coli ATP synthase catalytic complex (F1) (subunit composition 33). Diffraction data were collected to 3.15 Å resolution using synchrotron radiation. 

Use of a repetitive seeding protocol to obtain diffraction-quality crystals of a putative human D-xylulokinase. R. D. Bunker, J. M. J. dinkson, T. T. Caradoc-Davies, K. M. Loomes and E. N. Baker. Acta Cryst. (2012). F68, 1259-1262 [ doi:10.1107/S1744309112031181 ]. Synopsis: A putative human D-xylulokinase enzyme has been expressed, purified and crystallized using a repetitive seeding protocol. The trigonal crystals belonged to space group P31 or P32, with unit-cell parameters a = b = 101.87, c = 158.85 Å, and diffracted X-rays to at least 2.7 Å resolution. 

In situ X-ray data collection from highly sensitive crystals of Pseudomonas putida PtxS in complex with DNA. E. Pineda-Molina, A. Daddaoua, T. Krell, J. L. Ramos, J. M. García-Ruiz and J. A. Gavira. Acta Cryst. (2012). F68, 1307-1310 [ doi:10.1107/S1744309112028540 ]. Synopsis: The crystallization of both native P. putida transcriptional regulator PtxS and its complex with its DNA recognition sequence using the counter-diffusion method are reported. 

Ellman's reagent in promoting crystallization and structure determination of Anabaena CcbP. X.-X. Fan, Y.-F. Zhou, X. Liu, L.-F. Li and X.-D. Su. Acta Cryst. (2012). F68, 1409-1414 [ doi:10.1107/S1744309112034938 ]. Synopsis: Ellman's reagent oxidized the free sulfhydryl group of cysteine in Anabaena CcbP protein, facilitating its crystallization. 

Crystal sample pins and a storage cassette system compatible with the protein crystallography beamlines at both the Photon Factory and SPring-8. M. Fujihashi, M. Hiraki, G. Ueno, S. Baba, H. Murakami, M. Suzuki, N. Watanabe, I. Tanaka, A. Nakagawa, S. Wakatsuki, M. Yamamoto and K. Miki. J. Appl. Cryst. (2012). 45 [ doi:10.1107/S002188981203734X ]. Synopsis: A new cassette system for protein cryocrystallography has been developed. This system is compatible with crystal-exchange robots installed at both SPring-8 and the Photon Factory. 

A fast, simple and robust protocol for growing crystals in the lipidic cubic phase. M. Aherne, J. A. Lyons and M. Caffrey. J. Appl. Cryst. (2012). 45 [ doi:10.1107/S0021889812037880 ]. Synopsis: A simple robust manual protocol for producing crystals in the lipidic cubic phase in less than an hour is described. It is designed to provide newcomers to the in meso method for crystallizing membrane proteins with experience of preparing, handling and growing crystals in the sticky and viscous lipidic mesophase. 

Structural studies of human insulin cocrystallized with phenol or resorcinol via powder diffraction. F. Karavassili, A. E. Giannopoulou, E. Kotsiliti, L. Knight, M. Norrman, G. Schluckebier, L. Drube, A. N. Fitch, J. P. Wright and I. Margiolaki. Acta Cryst. (2012). D68, 1632-1641 [ doi:10.1107/S0907444912039339 ]. Synopsis: The effects of the ligands phenol and resorcinol on the crystallization of human insulin have been investigated as a function of pH. 

Measurement of detergent concentration using 2,6-dimethylphenol in membrane-protein crystallization. C. Prince and Z. Jia. Acta Cryst. (2012). D68, 1694-1696 [ doi:10.1107/S0907444912040176 ]. Synopsis: Membrane proteins require detergents for their stabilization in vitro, but detergents can interfere with downstream experiments such as crystallography. This work provides an efficient way to measure the concentrations of the glycosidic detergents most commonly used in crystallography. 

High Resolution Protein Crystals Using an Efficient Convection-Free Geometry. Alaa Adawy , Etienne Rebuffet , Susanna Tornroth-Horsefield , Willem J. DeGrip , Willem J. P. van Enckevort , and Elias Vlieg. Cryst. Growth Des., Just Accepted Manuscript. DOI: 10.1021/cg301497t. Publication Date (Web): December 5, 2012. Synopsis: Macromolecular crystallography is the most direct and accurate approach to determine the three-dimensional functional structure of biological macromolecules. The growth of high quality single crystals, yielding the highest X-ray resolution, remains a bottleneck in this methodology. Here we show that through a modification of the batch crystallization method, an entirely convection-free crys-tallization environment is achieved, which enhances the purity and crystallinity of protein crystals. This is accomplished by using an upside-down geometry, where crystals grow at the “ceiling” of a growth-cell completely filled with the crystallization solution. The “ceiling crystals” experience the same diffusion-limited conditions as in space microgravity experiments. The new method was tested on bovine insulin and two hen egg-white lysozyme polymorphs. In all cases, ceiling crystals diffracted X-rays to resolution limits beyond that for other methods using similar crystallization conditions. In addition, we demonstrate that the ceiling crystallization method leads to crystals with much lower impurity incorporation. 

Crystallization of the effector-binding domain of repressor DeoR from Bacillus subtilis. Jana Pisackova , Katerina Prochazkova , Milan Fabry , and Pavlina Rezacova. Cryst. Growth Des., Just Accepted Manuscript DOI: 10.1021/cg301551v Publication Date (Web): December 19, 2012. Synopsis: DeoR repressor of Bacillus subtilis negatively regulates expression of enzymes needed for metabolism of deoxyribonucleosides and deoxyribose. To gain structural information on regulation of DeoR function by small molecular ligand, we prepared crystals of C-terminal domain of DeoR in its free form and in complex with effector molecule deoxyribose-5-phosphate. To develop optimal procedure for crystallization, we have employed thermofluor assay as a tool for optimization of protein crystallizability. Monocrystals of three crystal forms were used to collect complete sets of diffraction data at synchrotron radiation source and will be used to.determine DeoR crystal structure. 

REACH: Robotic Equipment for Automated Crystal Harvesting using a six-axis robot arm and a micro-gripper. M. Y. Heidari Khajepour, X. Vernede, D. Cobessi, H. Lebrette, P. Rogues, M. Terrien, C. Berzin and J.-L. Ferrer. Acta Cryst. (2013). D69, 381-387 [ doi:10.1107/S0907444912048019 ]. Synopsis: A six-axis robot arm equipped with a micro-gripper is used to harvest protein crystals from their crystallization drops. Once harvested, the sample is inserted into the X-ray beam by the robot for direct data collection. 

Some practical guidelines for UV imaging in the protein crystallization laboratory. S. Desbois, S. A. Seabrook and J. Newman. Acta Cryst. (2013). F69, 201-208 [ doi:10.1107/S1744309112048634 ]. Synopsis: The use of UV imaging as a means of locating protein crystals is a fairly new tool, however not suitable for all protein crystallization trials. Practical examples of the strengths and some of the pitfalls of the technology are presented. 

Optimization of protein buffer cocktails using Thermofluor. L. Reinhard, H. Mayerhofer, A. Geerlof, J. Mueller-Dieckmann and M. S. Weiss. Acta Cryst. (2013). F69, 209-214 [ doi:10.1107/S1744309112051858 ]. Synopsis: The Thermofluor assay constitutes a quick and easy-to-perform high-throughput method with which it is possible to identify protein-stabilizing buffer compositions or small-molecule additives. 

A simple and effective calibration method to determine the accuracy of liquid-handling nano-dispenser devices. S. Rodríguez-Puente, J. Linacero-Blanco and A. Guasch. Acta Cryst. (2013). F69, 336-341 [ doi:10.1107/S1744309113000791 ]. Synopsis: A simple fluorescence-based calibration method that can be used to monitor the precision and accuracy of any liquid-handling nano-dispenser device is presented. 

A high-pressure cryocooling method for protein crystals and biological samples with reduced background X-ray scatter. C. U. Kim, J. L. Wierman, R. Gillilan, E. Lima and S. M. Gruner. J. Appl. Cryst. (2013). 46, 234-241 [ doi:10.1107/S0021889812045013 ]. Synopsis: A new crystal-hydration method has been developed for high-pressure cryocooling of protein crystals.

Towards protein-crystal centering using second-harmonic generation (SHG) microscopy. D. J. Kissick, C. M. Dettmar, M. Becker, A. M. Mulichak, V. Cherezov, S. L. Ginell, K. P. Battaile, L. J. Keefe, R. F. Fischetti and G. J. Simpson. Acta Cryst. (2013). D69, 843-851 [ doi:10.1107/S0907444913002746 ]. Synopsis: The potential of second-harmonic generation (SHG) microscopy for automated crystal centering to guide synchrotron X-ray diffraction of protein crystals has been explored. 

Imaging protein three-dimensional nanocrystals with cryo-EM. I. Nederlof, Y. W. Li, M. van Heel and J. P. Abrahams. Acta Cryst. (2013). D69, 852-859 [ doi:10.1107/S0907444913002734 ]. Synopsis: Electron imaging of three-dimensional protein nanocrystals using a Titan Krios electron microscope and a Falcon direct electron detector revealed structural details at higher than 3 Å resolution. Image processing with IMAGIC (cryo-EM single-particle analysis software) could improve the data to beyond 2 Å resolution. 

Effects of cryoprotectants on the structure and thermostability of the human carbonic anhydrase II-acetazolamide complex. M. Aggarwal, C. D. Boone, B. Kondeti, C. Tu, D. N. Silverman and R. McKenna. Acta Cryst. (2013). D69, 860-865 [ doi:10.1107/S0907444913002771 ]. Synopsis: Here, a case study of the effects of cryoprotectants on the kinetics of carbonic anhydrase II (CA II) and its inhibition by the clinically used inhibitor acetazolamide (AZM) is presented. 

Using high-throughput in situ plate screening to evaluate the effect of dehydration on protein crystals. A. Douangamath, P. Aller, P. Lukacik, J. Sanchez-Weatherby, I. Moraes and J. Brandao-Neto. Acta Cryst. (2013). D69, 920-923 [ doi:10.1107/S0907444913002412 ]. Synopsis: A novel high-throughput in situ plate-screening procedure is used to assess the effect of dehydration on a membrane-associated protein. In this case the dehydration improved the diffraction quality of the crystal. 

Integrated database of information from structural genomics experiments. Y. Asada, M. Sugahara, H. Mizutani, H. Naitow, T. Tanaka, Y. Matsuura, Y. Agari, A. Ebihara, A. Shinkai, S. Kuramitsu, S. Yokoyama, E. Kaminuma, N. Kobayashi, K. Nishikata, S. Shimoyama, T. Toyoda, T. Ishikawa and N. Kunishima. Acta Cryst. (2013). D69, 914-919 [ doi:10.1107/S0907444913001728 ]. Synopsis: Information from structural genomics experiments has been compiled and published as an integrated database.

Ben-Shem et al (Acta Cryst. (2003). D59, 1824-1827) in the crystallization of higher plant photosystem I found the detergent to chlorophyll ratio had to be carefully optimized in all purification steps in order to produce ordered crystals. Crystals were produced in 22.5 mM MES-bis-tris pH 6.6, 0.5% v/v PEG 400, 8.1 mM ammonium citrate, 6% w/v PEG 6000. Crystals appeared in 2 to 3 days and matured in size within two weeks time, yet the loss of sharp edges and a degradation of diffraction quality was observed after an additional two weeks time. The initial diffraction resolution of 20 Angstrom was improved to 6 Angstrom through a seemingly tedious refinement of isolation, crystallization and cryo conditions. Again, the detergent to chlorophyll ratio as well as the pea type and growing conditions with adjustment of the preparation to seasonal changes were essential to improvement of crystal quality.

When screening detergents as additives be sure to evaluate small amphophiles such as 1,2,3-heptanetriol, benzamidine, ethanol, dioxane, 1,6 hexanediol, ethylene glycol, and butyl ether for their ability to "manipulate" micelles.

Detergents can be used successfuly as crystallization reagents in oil based microbatch experiments. Measurements by Barenda et al and Loll et al indicated no significant loss of detergent will occur by migration of the detergent into the oil. 

References 
Thomas R.M. Barenda and Bauke W. Diskstra. Oils used in microbatch crystallization do not remove a detergent from the drops they cover. Acta Cryst. (2003). D59, 2345-2347. 
Loll et al. Compatibility of detergents with the microbatch-under-oil crystallization method. Acta Cryst. (2003). D59, 114-116.

Alkyl glucoside surfactants (detergents) can hydrolyze over time, producing an alcohol and a glucoside. This can sometime have an effect on the outcome of a crystallization experiment. When an older detergent solution produces crystals and a fresh detergent solution does not, try adding varying amounts of the alcohol to the fresh detergent solution to simulate the older detergent. 

For example, if an old MEGA-9 detergent produces crystals and a fresh MEGA-9 goes not, try adding varying amounts of nonyl alcohol to the fresh MEGA-9 solution to simulate a degraded MEGA-9 solution which would contan MEGA-9 and nonyl alcohol. For n-Octyl-ß-D-glucoside try adding octanol, and so on.

Under sitting or hanging vapor diffusion conditions where the protein-detergent complex crystallizes the free/bound detergent reaches equilibrium. When one opens the experiment the drop begins to evaporate and the detergent concentration will increase in the drop, which can dissolve or crack the crystals. This can also occur when stabilization/freezing buffers have too high a detergent concentration so consider this during cryo procedures. Some have reported 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. Consider that there is often constant detergent exchange between the solvent and the crystal except perhaps in instances where the detergent associated specifically with the protein in the crystal (Silver Bullet effect). One way to solve the problem of vapor diffusion set ups resulting in too high of detergent concentration after the drop is equilibrated is to set up the drops with lower initial detergent concentrations. As the concentrations of salt and PEG or other chemicals in the drop increases, the detergent CMC decreases, even for non-ionic detergents. Therefore, by dropping the detergent concentration, often to just below the apparent CMC, one may have a better chance at growing nice stable crystals.

A representative sample of the detergents offered by Hampton Research has been stability tested in solution and solid form. The detergents dodceyl-ß-D-maltoside, MEGA-9, octyl-ß-D-glucoside, CHAPS, CHAPSO, dodecyl-N-Ndimethylglycine, octyl-ß-D-galactoside, sodium cholate, and dodecyl-N-N-dimethylamine oxide were stored in solid form at 40 degrees Celsius for 7 days and at 25 degrees Celsius for 14 days. The detergent specifications for absorance, conductance, pH, percent fluorescence and purity by HPLC were measured before and after and all remained within specification. The detergents dodceyl-ß-D-maltoside, MEGA-9, octyl-ß-D-glucoside, CHAPS, CHAPSO, dodecyl-N-Ndimethylglycine, octyl-ß-D-galactoside, sodium cholate, and dodecyl-N-N-dimethylamine oxide in solution form were heated to 55 degrees Celsius for 2 hours and then retested for purity by conductance, absorbance and HPLC and all measurements showed no deterioration during the temperature cycle.

The following procedure for separating twins is reported in "Structural Basis for Double-Stranded RNA Processing by Dicer", Ian J. MacRae, Kaihong Zhou, Fei Li, Adrian Repic, Angela N. Brooks, W. Zacheus Cande, Paul D. Adams, Jennifer A. Doudna. Science 13 January 2006: Vol. 311. no. 5758, pp. 195 - 198. 

Crystals of the protein Dicer grown at 18°C by vapor diffusion in hanging drops composed of equal volumes of protein solution (9 mg/ml) and reservoir solution (28% PEG-400, 0.1 M MgCl2, 0.1 M NaCl, 5 mM TCEP, 1 mM DTT and 0.1 M MES, pH 6.5). Resulting crystals were pseudo-merhodrally twinned, producing an apparent spacegroup of P4222. Substituting MgCl2 with MnCl2 resulted in crystals that were macroscopically twinned. Individual crystals, which belong to the spacegroup P21212, were extracted by soaking the largest twinned crystals in reservoir solution containing 15% PEG-400 to weaken the twinning lattice contacts, followed by gentle mechanical prodding with a cat whisker. The resulting crystal shards were transferred back into full strength reservoir solution, cooled to 4°C for 1 hour and then cryo-cooled by plunging into liquid N2.

1) Try additives. 
2) Try DNA shuffling to introduce random mutations. 
3) Try agarose gel crystallisation. 
4) Try a new construct ("having tried everything else for 3 years before..."). 
5) Try to work on very small crystals and/or using a very small beam of a microfocus beamline at a synchrotron to isolate a single domain and get less twinning. 
6) Destabilise the crystal to separate the two halves (when possible) as in http://www.sciencemag.org/cgi/data/311/5758/195/DC1/1 
7) If the crystals are obtained with Mg exchange it to Mn. 
8) Change in crystallisation pH. 
9) Change in crystallisation temperature. 
10) You can try switching proteins. Lysozyme usually does the trick. 
11) Co-crystallisation with partner proteins/domains or with ligands. 
12) "Hmmm. Who knows, eh?" 
13) Change crystallisation setup (e.g., from microbatch to hanging drops). 
14) Test crystals at room temperature as well: "(EBV protease) which became twins only upon freezing, at room temperature the crystals were untwinned (but resolution was much worse as well)". 
15. See the following publication for an interesting method to resolve pseudo-merohedral twinning in SeMet crystals: 
Cansizoglu, A. E. and Chook, Y. M. (2007) Conformational heterogeneity of Karyopherinß2 is segmental. Structure, 15(11):1431-1441. 
"In an effort to obtain single selenomethionine Kapß2 crystals, mixtures of selenomethionine and native proteins were crystallized. A 
1:1 molar mixture of the two proteins gave single crystals..." 

Macromolecular crystal annealing (MCA) can help overcome increased mosaicity associated with cryocrystallography (Harp et al 1998, 1999). One might consider annealing if diffraction is uncharacteristically poor after flash cooling. The process cycles a flash cooled crystal to ambient temperature and then to cryogenic temperature and requires no special equipment. The annealing process does not improve a poorly diffracting crystal suffering from molecular disorder. The annealing protocol assumes that adequate cryo protection is available or that the crystal may be flash cooled using an oil and that the crystal diffracts well. The crystal is flash cooled a cryo stream. In the MCA procedure, the crystal is removed from the cryo stream and placed in a large (300 microliter) volume of the optimal reagent/cryo/oil solution from which the crystal was originally grown/mounted. Cover this drop to prevent evaporation. Incubation time at room temperature is typically 3 minutes. Extended incubation times are okay, but shortening the incubation time can produce inconsistent results. MCA has been successfully reported for both small and large crystals as well as for crystals with low (30%) or high solvent (70%) content. 

Other forms of crystal annealing, one termed flash annealing (Yeh and Hol 1998), the other termed annealing on the loop (Harp 1999), have been used successfully, especially on crystals with low (30%) solvent content. For flash annealing the crystals remain in the loop and on the mount. The cryo stream is diverted for 1.5-2.0 seconds, then reflash cooled for 6 seconds before repeating the process for a total of 3 rounds of rewarming and flash cooling. For annealing on the loop, a variable length of time for warming is used without multiple rounds of warming and reflash cooling. Warming time for cooling on the loop is determined by observing the crystal in the loop while the stream is diverted and waiting until the drop in the loop is clear before reflash cooling the crystal. Warming time is typically proportional to the size of the crystal. For the flash annealing and annealing on the loop methods one might carefully reduce the amount of liquid in the loop using a paper wick or micro wick as a variable to improve annealing results. But be careful not to let the drop dry too much. Finally, one might be prepared for the formation of ice on the crystal and loop when the stream is blocked. Ice can be removed with the delicate use of a paper wick or fiber. 

In closing, one may attempt the quick annealing methods first, although the MCA seems to be more general. 

References 
Macromolecular crystal annealing: evaluation of techniques and variables. Harp et al. Acta Cryst. (1999). D55, 1329-1334. 

Macromolecular crystal annealing: overcoming increased mosaicity associated with cryocrystallography. Harp et al. Acta Cryst. (1998). D54, 622-628. 

A flash-annealing technique to improve diffraction limits and lower mosaicity in crystals of glycerol kinase. Yeh and Hol. Acta Cryst. (1998). D54, 479-480. 

Small molecules which behave as electron and free radical scavengers can be used for co crystallization and soaking to reduce radiation damage to the macromolecular crystal. Recommended scavengers include oxidized glutathione, nicotinic acid, 5,5'-Dithiobis(2-nitrobenzoic acid) (DTNB), and ascorbic acid. Typical scavenger concentration is 0.2 M in the original crystallization reagent. A quick (less than 20 second) soak can be used to introduce the scavenger to the crystal. Prepare fresh solutions (add solid material to crystallization reagent in plate well) just before the soak. 

References 
How to avoid premature decay of your macromolecular crystal: a quick soak for long life. Brice Kauffmann, Manfreid Weiss, Victor Lamzin, and Andrea Schmidt. Structure 14, 1099-1105 July 2006. 

Investigation of possible free radical scavengers and metrics for radiation damage in protein cryo-crystallography. Murray and Garman. J. Synchrotron Radiation 9, 347-354, 2002. 

Blundell, T.L. and John, L.N. (1976) Protein Crystallography (New York, Academic Press). 

Here is the compiled 'tricks of the trade' for microbatch crystal mounting from the CCP4B community. 

Rebecca Page, Brown University, September 2006. 

Original Post to CCP4 community: 
I find mounting crystals grown in microbatch drops difficult compared to mounting crystals grown out of sitting drops, primarily because of the oil layer. Do most people simply use the oil layer as a cryoprotectant? If not, do you typically try to remove the oil prior to transferring the crystal to a cryoprotectant. I'd like to compile 'tricks of the trade' for mounting crystals out of microbatch plates. Any advice would be helpful and I'll compile the list of responses and repost for the community. 

SUMMARY OF RESPONSES 
Sorry if this sounds a crazy suggestion, but sometimes the simplest things work. Did you try to mount the crystal directly on a loop and see if it diffracts. The oil can be cryoprotectant. 
The setup would be: 
1- fish the crystal with a loop 
2-do not care if it cames across the oil layer and it retains the oil. 
3-mount in the cryostream 
4-shoot X-rays and see if you have diffraction 

****************** 

If everything fails and you have few more drops of crystals and do not know how to freeze mount, here is another way you can try :- If you have few crystals in the drop under oil and the drop size (excluding oil!!) is few microliters: add 10 microlitre of mother liquor (you can get the mother liquor conc. based on few trials; it should be few % more than the final conc. of the one you had used while setting up the microbatch, a decent start will be 5 % more ), allow it to stand for few minutes. Then use the classical cappillary mounting method to suck the crystals slowly out and onto a cover slip. If possible try to remove as much as possible (you may not be able to remove everything) the halo of oil surrounding the drop on cover slip (using the same capillary watching under microscope to make sure you are not sucking out the crystals). The use of mother liquor (the 10 ul) is to basically to reduce the oil that come along when you suck the crystals, so if necessary you can use more ul's. Now quickly scoop the crystal with a loop and dip in the cryoprotectant and freeze it, in the usual way. This has worked for me, though it may need some more standardization depending on your case. 

****************** 

In one case I had crystals grown under paraffin oil in microbatch, which required 25% glycerol for cryoprotection. However, when I transferred them directly to the base condition + 25% glycerol, they would invariably break up. So I ended up picking up a crystal with a loop, pulling it through the oil, and putting it on a coverslip. Then I removed the surrounding liquid and quickly replaced it with 4 microliters of base solution + 5% glycerol, observing the process under the microscope. The crystal seemed to hold up fine, and I then progressively replaced the solution with base condition + 10%, 15%, 20% and finally 25% glycerol. The resulting crystal froze nicely, and provided a 2A dataset. Any oil that was still around the crystal from the first transfer was probably completely removed by the cryo/washing steps, as other people have already described. 

****************** 

I think the success of mounting out of microbatch depends on the type of oil and plate you are using. We successfully are able to transfer crystals out of drops and into cryoprotectants using a standard hampton mounted cryoloop. You need a microscope that has a decent distance between the plate and actual lens apparatus so you can get the cryoloop down far enough into the well without covering up your viewpoint. When the drop contains the cryoprotectant already... then we directly transfer to liquid nitrogen and do not try and remove the oil.... at that point the oil only covers the solution which surrounds your crystal. One of the main benifits of vapour diffusion under oil vs. other sitting drop type methods is that the cry.

How does the age of a crystal affect X-Ray diffraction data? 

Reading and References: 

The crystal structure of yeast phenylalanine tRNA at 2.0 Å resolution: cleavage by Mg2+ in 15-year old crystals. Luca Jovinea,, Snezana Djordjevica and Daniela Rhodes. Journal of Molecular Biology, Volume 301, Issue 2, 11 August 2000, Pages 401-414. 

Abstract: We have re-determined the crystal structure of yeast tRNAPhe to 2.0 Å resolution using 15 year old crystals. The accuracy of the new structure, due both to higher resolution data and formerly unavailable refinement methods, consolidates the previous structural information, but also reveals novel details. In particular, the water structure around the tightly bound Mg2+ is now clearly resolved, and hence provides more accurate information on the geometry of the magnesium-binding sites and the role of water molecules in coordinating the metal ions to the tRNA. We have assigned a total of ten magnesium ions and identified a partly conserved geometry for high-affinity Mg2+ binding. In the electron density map there is also clear density for a spermine molecule binding in the major groove of the T?C arm and also contacting a symmetry-related tRNA molecule. Interestingly, we have also found that two specific regions of the tRNA in the crystals are partially cleaved. The sites of hydrolysis are within the D and anticodon loops in the vicinity of Mg2+. 

Atomic resolution structure of a succinimide intermediate in E.coli CheY. Simonovic M, Volz K. J Mol Biol. 2002 Sep 27;322(4):663-7. 

Abstract: Isomerization of aspartate to isoaspartate occurs spontaneously in proteins, causes changes in protein structures, and correlates positively with the aging processes of many organisms, including Alzheimer disease in humans. Aspartate isomerization proceeds through an unstable cyclic succinimide intermediate. There are few protein structure determinations that have characterized the intermediates and products of this isomerization reaction. Here we report the discovery of an unusually stabilized succinimide ring in the 1.1A structure of the Escherichia coli CheY protein, as determined from a crystal eight years old. The ring is formed by the side-chain of aspartate 75 and the backbone nitrogen of glycine 76 in an exposed loop of the molecule. Stabilization of the succinimide is through interaction of a sulfate ion oxygen atom with the imide nitrogen atom. Formation of the ring caused conformational changes in the loop, but did not alter the overall structure of the protein. 

Using rational screening and electron microscopy to optimize the crystallization of succinate:ubiquinone oxidoreductase from Escherichia coli. R. Horsefield, V. Yankovskaya, S. Törnroth, C. Luna-Chavez, E. Stambouli, J. Barber, B. Byrne, G. Cecchini and S. Iwata. Acta Crystallographica Section D, Biological Crystallography 
Volume 59, Part 3 (March 2003). 

Synopsis: Crystals of SQR that diffract to 2.6 Å were obtained by rational screening and sample quality analysis using electron microscopy. 

From the publication:"It proved critical to freeze the crystals within 72 h of crystallization set-up. Crystals that were frozen after this time limit showed no diffraction. This alteration in properties was apparent by the change in colour from deep orange to pale yellow that was observed in crystals more than four weeks old (Fig. 1c). The deep orange colour of the crystals is attributed to the presence of haem b within the protein. The loss or breakdown of haem, demonstrated by the change of colour in the crystals, could lead to structural instability and consequently loss of diffraction. "

We use a much easier test for mounting crystals at room temperature. Just coat the crystal with Paratone N (Hampton Research catalog number HR2-643) and mount your crystal in a standard cryoloop. The Paratone N will slow down evaporation enough - no special tools required. You don't even need to remove all the liquid as you would do for flash-cooling the crystal in Paratone N. And a major advantage is that you can use the same crystal to collect under cryo conditions and directly compare the impact of cooling the crystal. 

Reference: CCP4 Bulletin Board January 16, 2009 from Filip Van Petegem, The University of British Columbia.

Crystals can sometimes stick to the crystallization plate or slide. When this happens, try the following. 

1. If the crystal is stuck to a plastic plate or plastic slide, manipulate the plastic instead of the crystal. Using a needle, such as a Micro Needle, a Crystal Probe or syringe needle, stick the needle into the plastic a bit away from the crystal. Press hard and wiggle the needle enough to distort the plastic. The idea is to stretch and distort the shape of the plastic under the crystal and not touch the crystal. This movement in the plastic can free the crystal from the plastic. 

2. Change plastics. Crystals are reported to stick less to polypropylene and cyclic olefin copolymer (COC) plastics. The Micro-Bridge is available in polypropylene. Several sitting drop crystallization plates are available in COC, including Intelli-Plate and Swissci plates. 

3. Crystals grown in the presence of a gel. 0.1-0.2 % (w/v) agarose or silica hydrogel will grow inside the soft agarose gel. Therefore, they are mechanically protected and will not settle to the bottom of the sitting-drop well. When you harvest a crystal cut generously around it with a MicroTool, pick it up using a CryoLoop. 

4. Employ micro seeding for better control of nucleation. 

5. Carefully position a small piece of dry ice on the opposite side of the plastic from the crystal. The temperature change can sometimes free the crystal from the plastic plate. 

6. Sonicate the plate to free the crystals from the surface. 

7. Apply a very thin, smooth layer of vacuum grease or petroleum jelly to the plate or cover slide and set the drop on the grease.

Overlay the drop with perfluoropolyether cryo oil, paraffin oil, silicon oil or Parabar to prevent evaporation of the 2-propanol. Using a cryoloop, move the crystal into the oil, mounted the crystal and remove for storage or data collection. 

For cryo, try 5-30% MPD, or 5-30% Glycerol or dragging the mounting crystal through Perfluoropolyether Cryo Oil. 

Another option is to add a small amount of Perfluoropolyether Cryo Oil (HR2-814) to the crystallization drop before mounting the crystal. After adding the Perfluoropolyether Cryo Oil , mount the crystal using a Mounted Cryoloop. Withdraw the mounted crystal from the drop and the Perfluoropolyether Cryo Oil will coat the mounted crystal. Cryogenically cool the mounted crystal. 

Try growing the crystals under oil, and then soak the organic solvents into the drops, through the oil. This makes it much easier to harvest the crystals because the oil becomes saturated with the solvent and stops it from evaporating when you pick up the crystals. This approach can be used at the screening stage too. Try the Vapor Batch crystallization plate (DI-041)for this method. Reference: Mortuza, et al. High-resolution structure of a retroviral capsid hexameric amino-terminal domain. Nature 431 (2004), pp 481-485. 

For a hanging drop vapor diffusion experiment, remove the cover slide, invert and place on the microscope stage. Add several drops of the crystallization reagent, from the reagent well, to the glass slide, spotting the drops in a circular pattern about the drop contain the crystals. This local vapor barrier can slow or stop the movement of crystals for a minute or more, while the crystals are mounted. 

Providing temperature change will not damage the crystals, move the plate to a cold room and mount the crystals in the cold room. The cooler temperature can slow or stop crystal movement in the volatile drop.

The following is a recommendation for referencing nonperiodical documents on the internet such as the Hampton Research web site. Note: This material is extracted from the 5th edition of APA's Publication Manual (© 2001). 

For stand-alone document, no author identified, no date. 

If the author of a document is not identified, begin the reference with the title of the document. 

A general reference to the Hampton Research web site would be: 

Hampton Research web site. Retrieved April 27, 2018 from http://www.hamptonresearch.com 

If you are referencing the "Crystallization Tips A to Z" try: 

Hampton Research Crystallization tips A to Z. Retrieved April 27, 2018, from http://www.hamptonresearch.com/support/tips.php

If you are referencing a pdf from the Hampton Research web site: 

Hampton Research Crystal Screen Fundamentals. Retrieved April 27, 2018 from 
http://www.hamptonresearch.com/documents/product/0000000001-0000000643.pdf 

Thank you for referncing information retrieved from the Hampton Research web site.

Generally, chemicals must be disposed of in accordance with the local and national rules and regulations applicable to the site of use. Hampton Research material safety data sheets provide general information with regard to disposal. 

Care must be taken that only qualified personnel conversant with any possible hazards and all safety instructions be entrusted with the disposal or destruction of hazardous chemicals and materials. Always carry out work in an appropriate environment with the appropriate safety equipment (goggles, gloves, clothing, breathing equipment and facial mask). Be careful! 

For additional information see: 

"Prudent Practices in the Laboratory: Handling and Disposal of Chemicals", National Academic Press, Washington DC (1995), 412 pp. ISBN 0309052297 

"Hazardous Laboratory Chemicals, Disposal Guide", CRC Press, Inc., 2nd Edition, (1996) ISBN 1566701082 

"Destruction of Hazardous Chemicals in the Laboratory", Wiley-Interscience; 2 edition (March 1994) 520 pp. ISBN 047157399X

There are more than 1,600 research articles in the IUCr crystallography journals referencing Hampton Research crystallization research tools. 

Crystallization screens 
Structure of the Ebola VP35 interferon inhibitory domain. Daisy W. Leunga, Nathaniel D. Gindera, D. Bruce Fultona, Jay Nixb, Christopher F. Baslerc, Richard B. Honzatkoa, and Gaya K. Amarasinghea. PNAS, January 13, 2009, vol. 106. no. 2, 411–416. 

Crystal Screen and Crystal Screen 2 
The structure of a redundant enzyme: a second isoform of aspartate -semialdehyde dehydrogenase in Vibrio cholerae. R. E. Viola, X. Liu, J. F. Ohren and C. R. Faehnle. Acta Cryst. (2008). D64, 321-330. 

Hampton Research screens and kits 
Glycerol concentrations required for the successful vitrification of cocktail conditions in a high-throughput crystallization screen. R. Kempkes, E. Stofko, K. Lam and E. H. Snell. Acta Cryst. (2008). D64, 287-301. 

Crystal Screen 
Crystal packing of plant-type L-asparaginase from Escherichia coli. K. Michalska, D. Borek, A. Hernández-Santoyo and M. Jaskolski. Acta Cryst. (2008). D64, 309-320. 

Crystal Screen 
Structure of the Oncoprotein Gankyrin in Complex with S6 ATPase of the 26s Proteasome. Shigeyuki Yokoyama et al. Structure 15, 179-189, February 2007 

Ammonium sulfate 
Crystal structures of IRAK-4 kinase in complex with inhibitors: A serine/threonine kinase with tyrosine as a gatekeeper. Zhulun Wang et al. Structure 14, 1835-1844, December 2006. 

Natrix and Nucleic Acid Mini Screen 
Crystallization and preliminary X-ray diffraction analysis of an Escherichia coli tRNAGly acceptor-stem microhelix. C. Förster, M. Perbandt, A. B. E. Brauer, S. Brode, J. P. Fürste, C. Betzel and V. A. Erdmann. Acta Cryst. (2007). F63, 46-48. 

Crystal Screen 
Preliminary structural studies of the transcriptional regulator CmeR from Campylobacter jejuni. C.-C. Su, F. Shi, R. Gu, M. Li, G. McDermott, E. W. Yu and Q. Zhang. Acta Cryst. (2007). F63, 34-36. 

Crystal Screen, Crystal Screen 2, Grid Screen PEG 6000, PEG/Ion and Crystallization Plates 
Preliminary X-ray analysis of XC5848, a hypothetical ORFan protein with an Sm-like motif from Xanthomonas campestris. S.-K. Ruan, K.-H. Chin, H.-L. Shr, P.-C. Lyu, A.H.-J. Wang and S.-H. Chou. Acta Cryst. (2007). F63, 30-33. 

SaltRx Crystallization Screen 
Crystallization and preliminary crystallographic analysis of molybdenum-cofactor biosynthesis protein C from Thermus thermophilus. S. P. Kanaujia, C. V. Ranjani, J. Jeyakanthan, S. Baba, L. Chen, Z.-J. Liu, B.-C. Wang, M. Nishida, A. Ebihara, A. Shinkai, S. Kuramitsu, Y. Shiro, K. Sekar and S. Yokoyama. Acta Cryst. (2007). F63, 27-29. 

Hampton Research Crystallization Screens and CryoLoops 
Preliminary crystallographic analysis of avian infectious bronchitis virus main protease. J. Li, W. Shen, M. Liao and M. Bartlam. Acta Cryst. (2007). F63, 24-26. 

Crystal Screen, Crystal Screen 2, Grid Screen Ammonium Sulfate, Grid Screen PEG 6000, Additive Screen and StockOptions Salt 
Purification, crystallization and preliminary crystallographic characterization of the caspase-recruitment domain of human Nod1. T. Srimathi, S. L. Robbins, R. L. Dubas, J.-H. Seo and Y. C. Park. Acta Cryst. (2007). F63, 21-23. 


Hampton Research Crystallization Screens and Cryschem Plates 
Expression, purification and preliminary crystallographic studies on the catalytic region of the nonreceptor tyrosine kinase Fes. 
I. Gnemmi, C. Scotti, D. Cappelletti, P. L. Canonico, F. Condorelli and C. Rosano. Acta Cryst. (2007). F63, 18-20. 

Natrix 
Purification, crystallization and preliminary X-ray analysis of the BseCI DNA methyltransferase from Bacillus stearothermophilus in complex with its cognate DNA. Evangelia G. Kapetaniou, Dina 
Kotsifaki, Mary Providaki, Maria Rina, Vassilis Bouriotisa, and Michael Kokkinidis. Acta Cryst. (2007). F63, 12–14. 

Crystal Screen 
Structure of FliM provides insight into assembly of the switch complex in the bacterial flagella motor. Sang-Youn Park, Bryan Lowder, Alexandrine M. Bilwes, David F. Blair, and Brian R. Crane. PNAS, August 8, 2006 vol. 103 no. 32 11886–11891. 

PEG/Ion, Crystal Screen, Crystal Screen Cryo, and Cryschem Plate 
Crystallization and preliminary X-ray diffraction analys

Science must not impose any philosophy, any more than the telephone must tell us what to say. --G.K.Chesterton

When using stock solutions from the refrigerator or any stock solution that has been standing for some time, be sure to thoroughly mix the stock solution before use. Storing stock solutions in the refrigerator can result in condensation forming on the inside upper portion of the bottle. If this condensation falls into the stock solution it will create a heterogeneous stock concentration which in turn could make reproducing the experiment difficult. Condensation can occur with any sealed sealed when there is a temperature change in the environment. So it is possible for this phenomena to occur with room temperature stored stocks if the temperature in the room changes significantly. To avoid such problems it is generally good technique to mix crystallization reagents prior to pipetting.

Use a gel-loading pipet tip to dispense small drops with better control and accuracy of drop placement.

If you routinely mix the sample with the precipitant by aspirating and dispensing the drop several times try setting up the drop without mixing. Simply pipet the precipitant into the sample and let the two diffuse together without mixing. On the other hand, if you set up your drop without mixing, try mixing. In a survery during the 2005 RAMC meeting a show of hands survery among 100 participants showed most people do not mix their drops.

Communicate. Call (no e-mail allowed) a colleague ask them their favorite crystallization trick or tip. Offer up one or two of yours.

If you are using PEG 400 as a precipitant with some success try MPD. If you are using MPD try PEG 400.

Read the classics. Try Laboratory Experiments in Biological Chemistry by James B. Sumner and G. Fred Somers, Academic Press 1949.

Escape the local minima of using the same precipitants. If something worked once, great. Try it again but remember to try something new.

Try different temperatures. Room temperature and 4°C are the norm and work quite fine. Anything between 0°C and 37°C is fair game. If you do not have an incubator simply look for a cool or warm space in the building to incubate the plates.

Try free interface diffusion as a crystallization technique. This is easy to do and requires only a small amount of sample. Using a 20 microliter microcapillary pipet, pipet 2 to 5 ul of sample into one end of the capillary. Next, carefully pipet 2 to 5 microliters of precipitant SLOWLY into the capillary without introducing an air bubble. Seal the ends with wax or clay. It helps to pipet the less dense material onto the material of higher density (i.e. pipet sample onto 30% PEG 8000, pipet 15% isopropanol onto the sample). Use gel-loading pipet tips to load the sample and precipitant.

Tired of cutting filter paper for wicks to remove mother liquor from capillaries during crystal mounting? Try our paper wicks. The 55 mm X-fine work great in 0.75 mm capillaries.

Aging of polyethylene glygols (poly(oligo)-oxyethylene-based compounds) changes the chemical properties of common polyethylene glygols and detergents (Brij, Triton, C10E6, etc.), resulting in increased levels of aldehydes and carboxylates and peroxides to the system, but also by increasing metal binding and lowering pH. 

Aging is accelerated by light, warm temperatures and oxygen. To minimize the aging of polyethylene glycols store solutions in the dark or keep them covered. Freezing polyethylene glycol solutions slows the rate of aging much more so than refrigeration but refrigeration is preferred to room temperature storage for Use of amber or solid colored bottles or wrap clear bottles in foil. If you want to be really careful, purge them with argon and freeze them.

References 

A simple procedure for removing contaminating aldehydes and peroxides from aqueous solutions of polyethylene glycols and of nonionic detergents that are based on the polyoxyethylene linkage. William J. Ray, Jr. and Joseph M. Puvathingal. Analytical Biochemistry, Volume 146, Issue 2, 1 May 1985, Pages 307-312. 

Effect of chemical impurities in polyethylene glycol on macromolecular crystallization. Frances Jurnak. Journal of Crystal Growth, Volume 76, Issue 3, 2 August 1986, Pages 577-582. 

As a general rule, larger and finer crystals are obtained if the growth rate is minimized. If you are growing microcrystals reduction of their growth rate by more gradual equilibration with the reservoir may result in their attaining larger size.

Consider using a reducing agent as a variable in the crystallization. Include a small amount (0.001 to 0.01 M) of a mild reductant such as TCEP hydrochloride, cysteine, B-mercaptoethanol, glutathione, or dithiothreitol in the mother liquor. Beside protecting sensitive cysteine residues there is some indication that antioxidants play some other undefined role in crystallization. Try levels of reducing agent much higher than that generally thought necessary to maintain sulfhydryl function.

Once crystallization conditions are relatively well established, set up a crystallization trial and do not examine or disturb it in any way for several weeks. Premature handling of crystallization trials can convert a few promising growth centers into a massive shower of microcrystals.

Keep crystallization samples and hardware free from microbial presence by filtering all solutions through a 0.22 micron filter. Store stocks and working solutions in sterile containers. In most cases this negates the need for antimicrobials such as sodium azide.

Consider crystallization as a tool for bioseparation.

Proteins in high salt are usually more soluble at lower temperatures than higher temperatures. The reverse is generally true for PEG and MPD. Keep this in mind during initial screening and optimization. Keep in mind this is not always true.

pH adjustments in the drop can be made by adding a small amount of concentrated ammonium hydroxide to the drop or reservoir to increase the pH, or acetic acid to lower the pH.

If you are using organic solvents as precipitating agents be sure to include 4°C incubations with your trials. 

A 1:10 dilution can be defined as 1 part of stock plus 9 parts of diluent. For example, to make 1,000 microliters of 0.1 M buffer from 1.0 M buffer, simple pipet 100 microliters of 1.0 M buffer into a tube and add 900 microliters of diluent (water).

Percent Weight/Volume (% w/v) = the weight in grams of a solute per 100 milliliter of a solution. For example, a 50% w/v solution of PEG 6000 is made by adding exactly 50 grams of PEG 6000 solid to a total, final volume of 100 milliliters. NOT by adding 50 grams of PEG 6000 to 100 milliliters of water.

Urea and acrylamide contamination of RNA samples from denaturing gels cannot be removed by dialysis. Acrylamide and urea can be removed using a strong cation exchange resin. Joe Ng, UAB.

The path is as important as the endpoint. Vary the method, drop ratios, and reservoir volume to alter the path to the same endpoint as a variable during optimization.

Beware of flase positive inorganic salt crystals when using phosphate, carbonate, or borate buffers, especially in the presence of magnesiumm, calcium, zinc, and other metals. 

Magnesium in the presence of phosphate can form x shaped crystals. 

100 mM TCEP hydrochloride can form small hexagonal crystals in the presence of 50 mM Zinc acetate and 20% w/v Polyethylene glycol.

Stuck in a rut? Looking for new ideas? Dig through the NIST/CARB/NASA Biological Macromolecule Crystallization Database for fresh crystal growth ideas. Try http://bmcd.ibbr.umd.edu/.

Tired of condensation problems in the plates at 4°C? Fill two dummy plates with water, seal and place one at the top and the other at the bottom of your stack of plates. Voila! No condensation problem!

Polyethylene glycols are inherently unstable and prone to the presence and aldehydes, peroxides. The presence of these compounds increase with the age of the PEG and is promoted in the presence of UV light. One can slow the production of aldehydes and peroxide in PEG by storing the PEG at low temperatures and in the absence of light as well as oxygen (flood storage bottles with argon prior to long term storage).pH of the PEG will also vary with age as well as manufacture. As does the conductivity and ionic strength.

Never use sodium azide in crystallization reagents that contain heavy metals since the combination poses the danger of forming explosive metal azide salts (See Problems associated with the use of azide as an inhibitor of microbial activity in soil, M. Rozycki and R. Bartha, 1981, Appl. Env. Microbiol. 41: 833-836).

Recently we chatted about ways to alter the equilibration kinetics in a hanging or sitting drop vapor diffusion experiment. Another way to fool with the kinetics is perform the experiment at a cool temperature. Mikol et al have described how both the protein and precipitant are concentrated three to five time faster at 20°C than at 3°C. For details see V. Mikol, J.L. Rodeau, R. Giege, 1990, Anal. Biochem. 186, 332-339, Experimental determination of water equilibration rates in the hanging drop method of protein crystallization.

Always take into consideration the effect that salts, polymers, organic solvents, and additives might have on the pH of the buffer selected for the crystallization experiment. To be safe, check the pH of an experimental solution after all of the reagent components have been added, mixed, and the solution has equilibrated to the temperature where the buffer was titrated and the experiment is to be performed.

Although one typically uses 1,000 microliters of reservoir solution in a VDX or Linbro Plate, one can actually get away with 500 microliters of reservoir solution before one has to begin worrying about the excessive effects of evaporation which occurs when using polystyrene crystallization plates. The utilization of less volume in the reservoir can alter the kinetics of equilibration between the reservoir and the drop. Less volume equates to slower (longer) equilibration times. This is not necessarily a bad thing when screening for preliminary crystallization conditions. For a good read on the kinetics of macromolecular crystallization see "Kinetic Aspects of Macromolecular Crystallization" by Joe Luft and George DeTitta, page 110-130, Methods in Enzymology, Volume 276 Part A, 1997.

Flash-cooling crystals can sometimes increase the mosaicity of biological macromolecular crystals. In some case, macromolecular crystal annealing can reduce the mosaicity of flash-cooled crystals without affecting molecular structure. Crystal annealing involves cycling a flash-cooled crystal to room temperature and then back to cryogenic temperature. The procedure can also be applied to sometimes restore diffraction from flash-cooled crystals that were not properly handled to and from cryogenic storage. Crystal annealing does not seem to improve a poorly diffracting crystal suffering from molecular disorder. The essential features of the crystal annealing procedure are the following. The crystal is first flash-cooled. The crystal is removed from the cryostream and quickly transferred to a drop of cryoprotectant at crystal growth temperature and allowed to remain in the drop for at least three minutes. The drop should be covered to prevent evaporation. Finally the crystal is remounted on a loop and flash cooled. Reference: Macromolecular Crystal Annealing: Overcoming Increase Mosaicity Associated with Cryocrystallography, J.M. Harp, D.E. Timm, and G.J. Bunick, Acta Cryst. (1998) D54 622-628.

Soak crystals overnight in a solution comprised of the mother liquor utilized to grow the crystals plus 0.5 to 2% v/v glutaraldehyde. Glutaraldehye cross-links proteins through the epsilon amino group of lysines. Cross-linking can be used to stabilize fragile crystals for seeding or data collection. Reference: F.A Quiocho and F.M. Richards. Proc. Natl. Acad. Sci. U.S.A. 52, 833 (1964).

During the growth of a high quality crystal, identical components are incorporated into a periodic lattice. Hence sample homogeneity, the presence of uniform biological macromolecules is typically essential for the growth of high quality crystals.

If members of a homogeneous population alter their form then the population is no longer homogeneous. Stability of a population is important in order to maintain the homogeneity of the population. One should take measures to preserve the stability of the sample, and make the sample resistant to change in order to prevent denaturation and aggregation of the sample.

We set screens in anticipation of growing crystals. In reality we are looking for ways to alter the solubility of the sample in order to grow a crystal. Therefore, review screens looking for more than crystals. Look for variables that alter the sample's solubility. The big hitters are sample concentration, reagent type and concentration, pH, and temperature. 

Reverse vapor diffusion can sometime be used to crystallize a protein in low ionic strength. This is an alternative method to dialysis that is less tedious to set up but also allows the protein concentration to be become more dilute as the experiment proceeds, whereas dialysis can maintain a constant protein concentration. Begin with the concentrated protein in salt. The concentration of salt should be sufficient to maintain the solubility of the sample, perhaps 0.1 to 0.5 M salt. Set the sample for crystallization using the hanging or sitting drop vapor diffusion method. Pipet deionized water into the reagent reservoir. Pipet the high ionic strength protein sample onto the cover slide (hanging drop) or post (sitting drop). Do not add water to the sample. Using only deionized water in the reagent reservoir will allow water to diffuse from the reagent reservoir into the drop. As the drop size increases by vapor diffusion the ionic strength in the drop will decrease. The protein concentration will also decrease, so it is a good idea to begin with a concentrated protein sample. It is also helpful to avoid having too much salt in the samples since this may cause the sample drop to absorb too much water and dilute the protein concentration too much for crystallization. For cryo protection one can add glycerol to the reagent reservoir and mix this with the sample drop.

To crosslink fragile crystals for a chance at improved handling, solution changes or cryo try the following. If you have hanging drops then transfer the cover slide to a Cryschem 24 well sitting drop plate or place a Micro-Bridge into the 24 well hanging drop plate. Pipette 2 to 5 microliters of 25% glutaraldehyde into the sitting drop post and pipette your crystallization reagent into the surrounding reservoir. Seal the experiment. Allow the glutaraldehyde to vapor diffuse into the sample drop containing the crystal. 30 minutes to 6 hours is generally sufficient. Glutaraldehyde relies primarily on lysine residues so the number of lysines as well as temperature can be two significant variables influencing cross linking time. Note that amines will interfere with glutaraldehyde so ammonium sulfate or Tris buffer will have to be dialyzed or exchanged out and substituted before cross linking is attempted. 

Note: Glutaraldehyde is toxic and precaution was taken against exposure to the reagent by conducting the cross linking reactions in a chemical fume hood. 

Reference: J. Appl. Cryst. (1999). 32, 106-112 
A gentle vapor-diffusion technique for cross-linking of protein crystals for cryocrystallography. Carol J. Lusty.

Circle your drops with crystals or promising results. With a Cryschem plate sealed using tape, simply use a Sharpie to circle the drops that look promising and drops that have crystals. This makes it easy to spot these drops later during follow up. We suppose you could get fancy and use different color Sharpies to code the results, but a simple black Sharpie circle around a well is easy to spot, even through a stack of plates.

If you are able to obtain crystals of the native or wild type protein and cannot get crystals of variants or mutants of the native protein, try streak seeding. Streak seed from the wild type crystals into drops containing the variant/mutant. 

Glover et alstreak seeded from wild type crystals, either immediately or after a day of equilibration. The seeding stock solution was prepared by diluting a drop with small crystals 100-fold using the reservoir solution. Streak seeding was performed by dipping a hair into the seeding stock solution and rinsed twice in the reservoir. Crystals grew to 100-400 micron in 1 to 2 weeks. 

Reference: Principles of protein-DNA recognition revealed in the structural analysis of Ndt80-MSE DNA complexes. Jason S. Lamoureux and J.N. Mark Glover. Structure 14, 555-565, MArch 2006.

Jovine (2000) used an evaporation method to control excessive nucleation and produce larger crystals. 

A vapor diffusion experiment is set with the sample and reagent concentration where no crystals form when the drop and reservoir are in equilibration with one another. Nucleation is effected by opening each vapor diffusion experiment (remove cover slide, film or tape) for a precise amount of time. The amount of time used by Jovine was 100-120 seconds. The amount of time is empirically determined and will depend upon drop volume, reservoir volume, temperature, and reagent formulation. The opening increases the evaporation of water from the drop and creates a small and temporary increase in the relative supersaturation, hopefully enough to promote nucleation before the system returns to equilibrium when the experiment is sealed. If the drop remains clear, the procedure can be repeated until crystals start to appear. If too many crystals appear, reduce the amount of evaporation time. 

This method can also be applied to experiments which remain clear as a way to effect a change in the drop's relative supersaturation which could result in a crystal, precipitate or phase separation. 

Reference 
A simple technique to control macromolecular crystal nucleation efficiently using a standard vapour-diffusion set up. Luca Jovine. J. Appl. Cryst. Volume 33, Part 3, Number 2, 988-989 (2000).

Setting up vapor diffusion crystallization experiments against different reservoir solutions may have a profound effect on the outcome of crystallization experiments. It has been demonstrated shown that one way to increase crystallization space is to set the same conditions over different reservoirs (Newman 2005). So, rather than screen more reagents, one can screen the same reagent set over two or three different reservoirs (such as sodium chloride, ammonium sulfate and polyethylene glycol 3305) in addition to the crystallization reagent. Using the screen reagent in four sets of drops and then using the reagent in addition to three alternative reservoirs expands the crystallization space as the path of equilibration and endpoint is unique for each reservoir solution. 

Recommended reservoir solutions 
1.25 - 2.0 M Sodium chloride 
1.0 - 1.5 M Ammonium sulfate 
30-50% Polyethylene glycol 3350 
20% Polyethylene glycol 3350, 0.2 M sodium chloride 

Replacement reservoir solutions (approximate) 
0.5 M Lithium chloride = 35% PEG 4000 
1.5-2.0 M Lithium chloride = 2.5 M Ammonium sulfate 

Sodium chloride should be expected to act as a simple desiccant. Ammonium sulfate can desiccate and affect the pH of the drop through the gaseous NH3 which is in equilibrium with the NH4+ in solution. PEG 3350 should desiccate, although with different kinetics compared to the salts (Luft & DeTitta 1995). 

Alternative reservoir solutions have also been referred to as common dehydrants, a generic reservoir, and desiccants. The use of a single, simple, concentrated solution for the reservoir in vapor diffusion crystallization experiments has been happening for decades (Dunlop & Hazes, 2005; Hempel, 1968; Luft et al., 1994; McPherson, 1992). 

For 48 and 96 well crystallization plates set up manually, to speed delivery of the reservoir, the reservoir solution can be transferred to a multichannel pipetter basin (HR3-269) and delivered to the plate using a multichannel pipette. For automated experiments the reservoir solution can be transferred to a deep well block (HR3-105) or a multichannel pipetter basin (HR3-269). 

References 
Expanding screening space through the use of alternative reservoirs in vapor-diffusion experiments. Janet Newman. Acta Cryst. (2005). D61, 490-493. 

Single Crystals of Transfer RNA from Formylmethionine and Phenyalanine Transfer RNA's.Arnold Hempel et al. Science, New Series, Vol. 162, No. 3860 (Dec. 20, 1968), 1384-1387. 

A modified vapor-diffusion crystallization protocol that uses a common dehydrating agent. Dunlop & Hazes. Acta Cryst. (2005). D61, 1041-1048. 

Luft et al. (1994). J. Appl. Cryst. 27, 443-452. 

Luft & DeTitta. (1995). Acta Cryst. D51. 780-785. 

McPherson. (1992). J. Cryst. Growth, 122, 161-167.

Are you seeing skin form on the surface of your drops containing sample and reagent and wondering what to do? 

Skins on the drop are considered bad. These skins are believed to be a layer of denatured protein. Skins can slow down the rate of vapor diffusion. 

Pick away the skin before mounting your crystal. If the skin wraps around the crystal it can affect the limit of diffraction. 

To avoid skins one can try crystallize under oil(1). Either put a few microliters of oil on top of the sitting drop, or use the microbatch under oil method. 

Keep in mind that covering the sitting drop with oil will affect the rate of vapor diffusion equilibration. The following oils are ranked in order of the rate of equilibration allowed by the oil Paraffin Oil (slowest rate), Als' Oil (medium rate), Silicon Oil (highest rate). 

Reference: 
1. Pearl, B.O’Hara, R.Drew, S.Wilson. Crystal Structure of AmiC: the controller of transcription antitermination in the amidase operon of Pseudomonas earuginosa. EMBO Journal 13. (1994), pp 5810-5817.

Should one be concerned about vibration during crystallization? 

1. "Many years ago at Imperial College we did a fairly systematic experiment on vibration using lysozyme (hanging drops in Linbro trays). We placed one tray in the basement on a very solid concrete base, one tray was placed in an incubator, frequently used and known to vibrate a little (with frequent door opening and closing), and the third tray was placed on a 
rather strongly vibrating metal plate hanging from a ceiling on a piece of wire. The vibrations were produced by a small motor 
with an acentric piece of metal attached to its rotor. The trays were examined at regular intervals. Crystals grew in all of them. 
The basement tray gave fewer but bigger crystals, but this may have been a small temperature effect - temperature was not controlled very well. Crystals from all trays were nice, they diffracted to the same resolution, with virtually identical Wilson distribution. After this experiment we stopped worrying about vibration." Tadeusz J. Skarzynski, 2006 CCP4 Bulletin Board 

2. "...This is not to say, however, that absolute silence or stillness must prevail in the presence of growing crystals. At the Massachusetts Institute of Technology all the tRNA and protein crystals were grow in a cold room that had a giant compressor attached and contained, as well, several ancient centrifuges and shaker baths. One could observe standing waves in the reservoirs of all the vapor diffusion chambers and frequently could scarcely converse above the sound. Whether this had a positive or negative effect, we could not be sure. It is more likely that dramatic changes in the environment, as those caused by handling, are more disruptive than ambient conditions themselves." Alexander McPherson (1976) "The Growth and Preliminary Investigation..." in Methods of Biochemical Analysis, Volume 23, page 284. 

3. CCD Video Observation of Microgravity Crystallization of Lysozyme and Correlation with Accelerometer Data. E. H. Snell, T. J. Boggon, J. R. Helliwell, M. E. Moskowitz and A. NadarajahActa Cryst. (1997). D53, 747-755. 

Synopsis: Stepped growth rates and crystal movements have been observed by CCD video in microgravity lysozyme protein crystallization experiments which correlate with `g-jitter' accelerations, especially of low frequencies. The spurts and lulls of crystal growth may, therefore, explain the residual mosaic block structure seen in protein crystal mosaicity and topography measurements. 

Additional Comment by James Holton: They saw a very clear conneciton between vibrations and crystal growth rates and indeed crystal quality. Things like "astronaut exercising" 
coincided with increased crystal growth rate a short time later. So, it would appear that even in the most controlled environments vibration control can be a challenge, but at least on the Shuttle, where everything gets logged, you can (could) look for relationships.

And a comment from Eddie Snell: Basically, for vibration there are two factors, frequency and magnitude. Any high magnitude vibration is bad, i.e. the sudden short duration, hopefully low frequency one when you drop your tray :) For the same magnitude, low frequencies (a few Hz) are worse than higher frequencies (mains frequency). High frequencies are quickly damped in a liquid environment. Measuring vibrations is fairly easy, translating that measurement into something useful is not. In the case James mentioned, we were looking at the reduced acceleration environment. Certain crystallization processes lasted over a longer time period than on the ground - in this case the presence of a halo of depleted solution around the crystal caused by the diffusion limited transport of the protein to the crystal face. Every time an astronaut exercised, this halo broke down and a growth spurt occurred (except for one astronaut who didn't exercise very hard at all, however he/she shall remain nameless.

Structure of Mycobacterium tuberculosis mtFabD, a malonyl-CoA:acyl carrier protein transacylase (MCAT). Hemza Ghadbane, Alistair K.Brown, Laurent Kremer, Gurdyal S. Besraa and Klaus Futterera. Acta Cryst. (2007). F63, 831–835. The crystal structure of M. tuberculosis mtFabD, the mycobacterial MCAT, has been determined to 3.0 A ° resolution by multi-wavelength anomalous dispersion. Phasing was facilitated by Ni2+ ions bound to the 20-residue N-terminal affinity 
tag, which packed between the two independent copies of mtFabD.

Crystallizing ligands with limited or no water solubility can be tricky. Here are some suggestions. 

A review on the crystallization of protein ligand complexes can be found in Crystallization of protein–ligand complexes, Annie Hassel et al, Acta Cryst. (2007). D63, 72–79. 

Vary the ratio of protein to ligand Try 1:3, 1:5 or 1:10 ratios of protein:ligand. This can make a huge difference. There are instances where the organic solvents (DMSO, Ethanol) used to dissolve the ligands interfere with the crystallization process. In such instances, try adding the dry compound to the crystallization drop with the protein. Or add the dry compound to the protein solution and put on a shaker overnight with gentle rotation. Centrifuge the next morning and set up your crystallization screens. It does not matter if it does not appear if the solid compound dissolved. Enough solid compound may go into solution to effect ligand binding. 

Successful complexation depends on the concentration of protein, ligand, and the Kd of the protein-ligand complex. For Kd much greater than protein concentration, try the ligand concentration at greater than 10 x Kd. As Kd approaches the protein concentration, roughly superstoichiometric quantities will be sufficient for full occupancy. For Kd less than protein concentration, stoichiometric quantities of ligand will typically suffice. In general, the ligand concentration should be such that it is near saturation on the binding isotherm. 

Solubilize the ligand in DMSO so it is maximally concentrated (100mM works fine). Add enough ligand to achieve two to three fold excess ligand to protein. Keep the DMSO concentration to no more than 3% to avoid damaging the protein. Set crystallization experiment using this sample. If you cannot achieve a high enough stock concentration of DMSO to be below the 3% threshold, dilute you protein in the storage buffer to approximately 1mg/ml. Add compound to two to three fold excess, incubate and co-concentrate to the desired concentration. This may help to avoid the DMSO shock. Alternatively one can incubate the concentrated protein with the compound solubilized in water for 24 to 48 hours and solubility of the ligand will be sufficient to complex with the protein. (Carsten Schubert ccp4bb December 2007) 

One method that worked for me was to dissolve my ligand in 100% DMSO, as suggested in the previous response, then add a 3 molar excess of ligand to protein so that the final concentration of DMSO in the protein-ligand solution was no greater than 10% - of course the maximum concentration of DMSO that your protein can suffer will be protein specific but you could investigate this by incrementally adding DMSO to just a solution of your protein at your working crystallization concentration then measuring scattering at 600nm in a spectrophotometer to determine the critical DMSO concentration that causes your protein to precipitate (if you have sufficient protein to 'waste'!). If your ligand binds tightly to your protein at an equimolar concentration you can then remove excess ligand and DMSO by passing your sample through a G25 Sephadex column. (Rob Hussey ccp4bb December 2007) 

If you grow crystals in polyethylene glycols or similar reagent you might try to solubilize the compound in a small amount of this reagent. This is helpful if you want just a 1:4 protein:ligand ratio. Sometimes solubility is low even in 5% DMSO (or diluted solutions of glycerol, alcohols and similar molecules). In these cases setting up drops in the presence a saturated solution and some precipitate of the compound may also lead to good co-crystals. As one molecule passes from the saturated solution to the “bound” state, a new molecule is solubilized from the precipitate, which gradually dissolves and passes from the solution to its binding site in the protein. Indeed, this sounds like a “soaking” experiment and it works well if the compound is colored, so that you can see if the crystals actually become of the same color. Just remember to wash them thoroughly before measurements, in order to remove traces of the ligand precipitate that would result in poor diffraction. (Marco Mazzorana ccp4bb December 2007) 

We routinely obtain structures from protein solutions with a big pellet of ligand in the bottom of the tube. For co-crystallizations we add 1mM compound to a 0.3mM solution of the protein and incubate overnight. Many of the compounds are only soluble to 50 micromolar, so we get a lot of precipitate. The next day, we spin the tube at high speed, and use the supernatant for crystallization trials. We have started from 100 mM stocks in 100% DMSO or ethanol. This has worked for compounds ranging for picomolar to micromolar affinity, which surprised us, but it worked. (Kendall Nettles ccp4bb December 2007) 

We normally prepare our ligand stocks in DMSO and add this to the protein in 3-fold molar excess. The majority of our ligands are quite insoluble and precipitate when the DMSO concentration decreases upon addition to the protein……. so I am not surprised that you are seeing this. If your compound does not bind your protein tightly, you might consider using a 5-fold molar excess of ligand. 

Some proteins crash out if the protein concentration in high when you add the ligand. For those situations, we complex the ligand with dilute protein (1-2 mg/ml), and then concentrate this for crystallization trials. I have had proteins where we had to complex the dilute protein with ligand, and then let it sit overnight at 4 degrees Celsius before we concentrated the protein. We normally incubate the protein+ligand at 4 degrees Celsius for 1-3 hours for binding before we set up the crystallization experiments. Another scenario might be addition of ligand to the protein followed by incubation at room temp for about 1 hour. Then centrifuge at 4 degrees Celsius, keep protein at 4 degrees Celsius and set up your trays. 
(Annie Hassell ccp4bb August 2011) 

Try solubilizing the ligand in low molecular polyethylene glycol (PEG 200, PEG 300 or PEG 400). 

When adding ligands to a drop containing crystal, consider optimizing for smaller crystals. 10-20 micron crystals can reach 80% occupancy in less than 30 seconds. A short soak time can minimize or prevent crystal damage. Larger crystals can equal longer soak times can equal higher risk for crystal damage. 

Recommended reading 
Guidelines for the successful generation of protein–ligand complex crystals. Ilka Muller, ActaCryst.(2017). D73, 79–92.

Glutaraldehyde to Differentiate Salt from Protein Crystals 
Place the crystals in a low ionic strength buffered solution containing 0.2-.2.0% v/v glutaraldehyde. Protein crystals will quickly be fixed quickly (faster than they dissolve) into a light golden, gelatinous lump. Sometimes the crystals retain a crystal-like shape, other times the crystals leave just a rubbery blob. In contrast to protein crystals, salt crystals should dissolve over time and should not be colored. 

One can add a small amount of 2% v/v glutaraldehyde to the protein drop. Protein crystals in the presence of glutaraldehyde will then turn a light golden color. Crystals fixed like this can then be put into a low ionic strength solution where salt crystals should dissolve. You can easily transfer glutaraldehyde into a protein drop by vapor diffusion by adding glutaraldehyde to the reservoir to make it 2-3% v/v glutaraldehyde. 

Or, one can add enough glutaraldehyde into the reservoir to make it 0.5-1.0% glutaraldehyde. A nice side effect of using the reservoir is that the reservoir is now a control. If the reservoir turns yellow it means you have free amines such as Tris or ammonium and this could interfere with the crosslinking of the protein. 

A caveat to using this method is that there should be no free amines around other than on the protein (i.e., no ethanolamine or Tris buffer, no ammonium ions, etc.). 

Another caveat, the yellow color, which is due to Schiff's base formation, is harder to see in warm halogen light when you are looking at small or thin crystals. Using a blue filter between the light source and the sample can remove the yellow from the light source. Or use cool, white lights such as LED light. 

Finally, keep some buffer around that is suitable for solubilizing the protein. If you are not sure about the color change, just add buffer to the crystals and watch if they dissolve. If they don't dissolve when treated with glutaraldehyde, they are protein crystals. If they dissolve, they are salt crystals. 

Note: Glutaraldehyde is toxic, causing severe eye, nose, throat and lung irritation, along with headaches, drowsiness and dizziness. Glutaraldehyde is quite volatile with a strong, obnoxious odor. 

A suggestion from Michael Garavito, Michigan State University.

1. Screen additives (See Hampton Research Additive Screen and Detergent Screen). 

2. Use a crystallization screen as an additive screen. Make the reservoir 90% original reagent plus 10% crystallization screen reagent (Crystal Screen, Index, etc). Repeat using the entire screen. 

3. Change the crystallization method. Sitting drop, hanging drop, microbatch, dialysis, free interface diffusion... 

4. Try limited proteolysis. Reference: In situ proteolysis for protein crystallization and structure determination. Nature Methods - 4, 1019 - 1021 (2007). 

5. Add fresh protein to the drop once crystal growth has ceased. 

6. Add water to the drop once crystal growth has ceased. Crystals may dissolve and grow larger or a new crystal form. 

7. Try the Silver Bullets screen from Hampton Research. Reference: Searching for silver bullets: An alternative strategy for crystallizing macromolecules. Alexander McPherson and Bob Cudney. Journal of Structural Biology 156 (2006) 387-406.

A set of four colored Sharpie pens can function as a simple scoring system for crystallization experiments. Use a black Sharpie to label your plate, a green to indicate crystals, a red for precipitate and a blue for unknown or curious result. An unmarked well indicates a clear drop. When scoring 96 well plates put a color dot over the reagent well. Such reagent wells can easily accept 3 different dots, so if the results change over time, one can add different colored dots, starting with the first scoring on top and then subsequent dots below as needed. When scoring a Cryschem plate, circle the reservoir using the same color scheme. Make a quarter of a circle about the reservoir with your first score, so if the results change over time, so can the score as you complete the circle. Plates are often scored immediately after set up, after 24 hours, at one week and then two weeks. To review your scoring, place the scored plate on a piece of white paper to look for hits and trends based on color. Retain plates until the drops dry out or salt crystals form in the reservoir. 

Sharpie is a registered trademark SANFORD or its Affiliates. 

When greasing crystallization plates, it is often useful to leave a small gap in the bead of grease applied to the plate. For example, begin greasing at 12:00 and apply the bead clockwise until 10:00. A gap with no grease is left between 10:00 and 12:00. Place the cover slide onto the bead of grease. Depress the slide onto the grease. As the slide is pressed onto the bead of grease, pressure is allowed to escape through the gap. Before the slide bottoms out on the plate, twist the slide approximately 15 minutes (90°) to smear the grease across the gap and properly seal the cover slide to the plate.

Crystallization by interface diffusion was popularized by Ray Salemme (F.R. Salemme 1972, Arch. Biochem. Biophys. 151, 533). The method is also called free interface diffusion and liquid-liquid diffusion. The methodology is straightforward. Interface diffusion is classically performed in a cylinder with a small diameter, such as a capillary. Using a capillary with both ends open, the sample is pipetted into the capillary. Next, the crystallization reagent is added to the capillary. For true interface diffusion to occur, the sample and the reagent should touch one another inside the capillary. This can be accomplished by making sure the sample is positioned at the inside end of the capillary before adding the crystallization reagent. When adding the crystallization reagent, use a pipet tip with a small diameter opening such as a gel loading tip. Carefully and slowly pipet the reagent from the pipet tip and allow a bead of the reagent to form at the tip. Hold the capillary parallel to the ground and gently touch the tip of the capillary to the reagent bead. With a combination of gently pipetting, capillary action and proper capillary tilt, fill the capillary with the appropriate amount of crystallization reagent without air gaps. Done correctly, the inside of the capillary should contain a single cylinder of liquid. Filling the capillary should be done gently and slowly to minimize mixing the sample and the reagent. Actually, one should add the more dense solution into the capillary first, followed by the less dense solution to minimize mixing. If done properly, one can see an interface between the sample and the reagent. It might be best to start with equal amounts of sample and reagent. Later, one can vary the ratio of sample to reagent for optimization. Once filled with reagent and sample, the capillary can be sealed with a non-drying clay or wax (sealant). Be careful not to add too much sealant at a time to the end of the capillary since the sealant will displace the liquid in the capillary and too much sealant will result in liquid being lost from the capillary. Storage of the capillary can be done in several ways. Some like to simply stick the capillaries vertically into clay or wax. But this means one must remove and contort the capillary for viewing under a microscope. Instead, one can use a petri dish (square ones work really well) and line the petri dish with two thin lines of wax or clay about the size of a toothpick. The lines of wax or clay should be parallel to one another and positioned apart from one another just under the length of the capillary. The capillaries can be set upon the lines of clay in neat rows. A dozen or so capillaries can be stored in a small square petri dish. Storage and viewing is quick and simple. The petri dish also facilitates easy labeling. 

A nice feature of interface diffusion crystallization is the potential of a concentration gradient of sample and reagent along the length of the capillary. Although pipetting, gravity, and movement make this interface and gradient less than perfect, gradients do form and one can often visualize the concentration gradient as crystals of different size and number form at different points along the gradient (length of capillary), presumably due to the change in relative supersaturation along the length of the capillary. For example, numerous, small crystals might form at high levels of supersaturation, fewer and larger crystals might form at ideal levels of supersaturation, and no crystals may form at very low levels of supersaturation. 
Permutations of interface diffusion include the following. Doing it in microgravity. Freezing one layer before adding another to ease pipetting (good for larger volumes). Freezing the entire experiment then slowly melting the slug to introduce temperature as a crystallization variable. Placing a gel plug between the sample and the reagent. Using high throughput plates with tall, thin cylinder-like wells for “vertical”

J is for Jeffamine® Reagent 

The Jeffamine polyoxyalkyleneamines contain primary amino groups attached to the terminus of a polyether backbone. They are thus “polyether amines.” The polyether backbone is based either on propylene oxide (PO), ethylene oxide (EO), or mixed EO/PO. Jeffamines are synthesized as either monoamines (M-series), diamines (D series), or triamines, and are made in a variety of molecular weights, ranging up to 5,000. The ED-series are aliphatic diamines structurally derived from the propylene oxide capped polyethylene glycol. 
The wide range of molecular weights, amine functionality, oxide type, and distribution provides flexibility in synthetic design of compounds made from Jeffamine Reagents. For the most part, Jeffamine Reagent products undergo typical amine reactions and are low viscosity liquids, exhibiting low vapor pressure. 
Jeffamine® Reagents originated chemically at the Texaco Chemical Company as lubricants and fuel additives and are now most frequently used in manufacturing adhesives, coatings, epoxies, and curing agents. Scary what we crystal growers will use as crystallization reagents, is it not? 
Jeffamine Reagents worked their way into the protein crystallization community in the late 70’s when Dulio Cascio (then in the lab of Alexander McPherson at the University of California Riverside now University of California Los Angeles) evaluated them as polymeric precipitants “similar” to the polyethylene glycols. 

Alexander McPherson published the use of Jeffamine Reagents in a crystallization publication “Two approaches to the rapid screening of crystallization conditions” J Crystal Growth 1992, 122: 161-167. 
The first protein structure solved using Jeffamine Reagents was that of Xylose Isomerase in the publication by Lloyd et al, “Crystallization and preliminary X-ray diffraction studies of xylose isomerase from Thermoanaerobacterium thermo sulfurigenes strain 4B” J Mol Biol 1994, 240: 504-506. 
Jeffamine Reagents are discussed by Lesley Lloyd Haire and others in Terese Bergfor’s wonderful book “Protein Crystallization - Techniques, Strategies, and Tips, A Laboratory Manual (IUL Biotechnology Series 1999 ISBN 98-075232). 
The following Jeffamine Reagents have been tried in protein crystallization experiments: 

• Jeffamine D-230 Reagents 
• Jeffamine D-400 Reagents 
• Jeffamine ED-600 Reagents 
• Jeffamine ED-900 Reagents 
• Jeffamine ED-2001 Reagents 
• Jeffamine M-600 Reagents 

Hampton Research has experienced the most success with Jeffamine ED-2001 Reagent and Jeffamine M-600 Reagent. 

Jeffamine Reagents can be used “like polyethylene glycols” in the crystallization of proteins, peptides, and nucleic acids. In fact, some crystal growers substitute Jeffamine Reagents for PEGs during optimization to see if using them can improve the crystals or offer new conditions for further optimization. 
Jeffamine Reagents can be formulated with most of the salts, buffers, organic solvents, and additives used in biological macromolecular crystallization. 

There is very little literature describing the use of Jeffamine Reagents as crystallization reagents and not a single report of them being used as a crystallization reagent in the Biological Macromolecular Crystallization Database at the time of this writing. 
(http://wwwbmcd.nist.gov:8080/bmcd/bmcd.html) 
Obtaining crystals in Jeffamine Reagents is a mixed bag. It is a good thing since there are crystals. It is a bad thing since they are a pain to formulate. 

The formulation of Jeffamine Reagents as crystallization reagents is tricky and tedious. They are very alkaline chemicals and must be titrated to neutrality or the desired experimental pH before use. Titration is typically performed using hydrochloric acid. A significant amount of hydrochloric acid must be used to titrate the Jeffamine Reagents to pH values between 2 and 11. pH changes are slow until pH 9, and change very rapidly as one approaches neutrality. The addition of hydrochloric acid results in a significant temperature increase in the reagent. Accurate pH recordings require repeated pH, cool, pH, titrate; repeat cycles. Many salts, including anions and cations, can be used in conjunction with Jeffamine Reagents. Typical salt/Jeffamine Reagent ratios and concentrations are similar to those used with “related” polyethylene glycols. Phase separation can be a bigger problem with Jeffamine Reagent / salt mixtures than for PEGs, but can be minimized by titrating the Jeffamine Reagent to neutrality or close to the final working pH before adding the salt and/or buffer. Having as much water present in the formulation also helps prevent phase separation. 
Jeffamine M-600 Reagent is utilized as a crystallization reagent in Crystal Screen 2 (Catalog Number HR2-112). 

Hampton Research offers the following preformulated, ready-to-use Jeffamine Reagents: 
Jeffamine ED-2001 Reagent 50% w/v solution pH 7.0, 200ml sterile filtered (HR2-597) 
Jeffamine M-600 Reagent 50% w/v solution pH 7.0, 200ml sterile filtered (HR2-501) 

Hampton Research also offers the following Jeffamine Reagent which has not been titrated: 
Jeffamine M-600 Reagent 100% solution pH >12, 200 ml (HR2-503) 

Jeffamine is a registered trademark of the Huntsman Petrochemical Corporation.

Kosmotropes (order-making) or lyotropes are ions that display strong interactions with water and generally stabilize proteins. This stabilization effect is brought about by increasing the order of water and increasing its surface tension.

Examples of kosmotropes include, but are not limited to, sodium citrate, sodium sulfate, trimethylamine N-oxide, proline, ectoine, trehalose, glycine betaine, 3-dimethylsulfoniopropionate, ammonium sulfate, lithium sulfate, sodium phosphate, and magnesium chloride.

Named from the Greek kryptos or “hidden”, krypton is neither green, nor a solid material that can defeat Superman. Rather it is another noble gas discovered in 1898 by Ramsay and Travers. It ranks sixth in abundance in the atmosphere. Krypton gas is used in various kinds of lights, from small, bright, flashlight bulbs to special strobe lights for airport runways. As with the other noble gases, krypton is isolated from the air by liquefaction. 

Krypton and crystals. The use of xenon and krypton at high pressure is becoming a popular method for the phasing of proteins. There are several homemade as well as commercially available pressure cells for creating xenon and krypton derivatives. 

In essence, the crystal is exposed to the krypton or xenon gas at high pressures for a short period of time in order to allow the krypton or xenon to bind to the protein within the crystal. Following incubation and depressurization, the crystal is flash cooled in liquid nitrogen and mounted onto a goniometer in a cryostream for X-ray diffraction analysis. Freezing of the crystal is necessary to prevent the diffusion, release and loss of gaseous krypton or xenon. As a side note, methods of generating and analyzing xenon derivatives in glass or quartz capillaries have been described (see references below). 

Krypton and xenon binding sites are generally different from those for other heavy atom sites so the screening of krypton and xenon can be used as a follow up when the initial soaks in heavy atoms are not successful. 

When the specialized hardware is available for screening krypton and xenon under high pressure, the method is a convenient and rapid way for screening successful derivatives for phasing. 

References: 
• High-pressure krypton gas and statistical heavy-atom refinement: a successful combination of tools for macromolecular structure determination. Schiltz, M., Shepard, W., Fourme, R., Prange, T., de La Fortelle, E. and Bricogne, G. Acta Cryst. D53: 78-92, 1997 
• Exploring hydrophobic cavities in proteins using xenon and krypton noble gas. Prange,T., Schiltz, M., Pernot, L., Colloc’h, N., Longhi, S., Bourguet, W. and Fourme, R. Protein Struct. Funct. and Genetics 30(1): 61-73, 1998. 
• Solubility of krypton and xenon in blood, protein solutions, and tissue homogenates. Yeh, S.Y., Peterson, R.E. J Appl Physiol 5: 1041-1047, 1965. 
• Flash freezing isomorphous xenon or krypton derivatives of protein crystals. Sauer, O., Dutzler, R and Kratky, C. ECM 17, Seventeenth European Crystallographic Meeting (Lisboa, Portugal 24/28 aug.1997). Book of Abstracts p 18 (ref. MS1.6-4) 
• Xenon and Krypton at LURE http://www.lure.u-psud.fr/sections/Xenon/XENON.HTM 
• Tutorial for Krypton-Elastase SIRAS refinement http://utica.med.jhmi.edu/sharp/tutorials/KrEl.html

Sialidase I from Clostridium perfringens (Glyko, Inc. Rosedale, NY, USA) was used by Ogawa et al (Acta Cryst. (2003). D59, 1831-1833) to remove terminal sialic acid residues and minimize the heterogeneity of the glycosyl structure on the protein and enhance crystallization. The protocol utilized 1 mU of Sialidase I in 50 microliters of 50 mM Tris-HCl buffer pH 6.8 at 310 K for 12 hours.

Xing and Xu (Acta Cryst. (2003) D59, 1816, 1818) recently describe the crystallization of the PX domain of cytokine independent survival kinase in ammonium sulfate. Optimization of the crystallization seemingly involved the subtle manipulation of several crystallization variables, including using drop ratios (1:2 drop ratio), the removal of DTT to allow crystal growth after nucleation, as well as the substitution of sodium malonate for ammonium sulfate. The authors reported the sodium malonate dramatically increased the reproducibility of the crystals and also acted as a good cryoprotectant. Crystals grown from 2.0 M sodium malonate could be cryo protected directly from the drop. The crystals grown in sodium malonate were more resistant to physical shock compared to those grown in ammonium sulfate.

If one is able to obtain crystals of a protein but the crystals do not diffract well, try further purification of the sample and repeat the crystallization conditions. Often time crystals do not diffract well because of impurities or aggregates present in the sample. Impurities in the sample can cause dislocations and defects in the crystal, which can lead to poor diffraction. Further purification can sometimes remove these impurities and aggregates, resulting in crystals with improved diffraction. Often times the reason crystals do not diffract can be traced to the molecules in the crystal not being well ordered in the lattice so there is conformational flexibility or the molecules do not take a desirable or fixed conformation because the lattice forces are too weak. Or sometimes there are severe defects in the crystal lattice, which lead to a disorientation of the crystal lattice.

Try increasing the drop size to grow larger crystals.

Centrifuge the sample at 2,000 g for 10 to 15 minutes immediately prior to set up to remove aggregates and amorphous material.

During optimization of crystallization conditions, examine any additive that might for some reason tend to stabilize or engender conformity by specific interaction with the macromolecule.

To reproduce crystallization conditions from a screen without taking additional solution from the screening kit (and making all your lab partners want to do evil things to you) simply set an additional drop or drops on the cover slide next to the original drop. One can set multiple drops on the same cover slide over the same reservoir. Pipet the sample onto the cover slide, then pipet the precipitant from the reservoir into the sample drop. Mix and seal the cover slide over the reservoir.

It has been suggested that lattice contacts within a crystal can be manipulated using ionic strength which in turn can be used to influence crystal morphology. Rather than the type of salt used (sodium chloride, sodium acetate, sodium citrate, etc) one may manipulate the concentration of the salt ot change crystal morphology. Typical salt concentrations to be screened for such manipulation is 0 to 500 mM.

Explore as many optimization opportunities for crystallization as possible. Optimize every hit from a screen. When you reach an optimization dead end for a particular hit, review the screens again and optimize crystal leads from other hits. Optimize different hits simulatanously if sample availability permits. This can save time if one hit leads a a dead end (poorly diffracting crystal, non-isomorphous derivative, etc).

One's tendency is to optimize the largest, best looking crystals. Common sense. However, there are plenty of reports where small, miserable, ugly crystals have eventually produced the best data, so do not ignore the ugly ducklings.

Control is a good thing. Having a handle on the variables which affect the crystallization and the ability to control these variables is key to the reproducible growth of high quality crystals.

When working with ADP, ATP, NAD, NADH or other chemically unstable ligands as crystallization additives, be sure to use freshly prepared working solutions or properly stored stocks. These agents are sensitive to degradation over time which can influence the outcome of a crystallization experiment. Also, consider the pH of these ligands in solution since the pH of the additive agent can influence the final pH of the drop. It might be necessary to titrate the ligand solution before addition to the drop.

How might one increase the size of a crystal which is grown from dialyzing the protein into pure deionized water? The following replies were posted on the cc04bb (http://www.ccp4.ac.uk) on October 21, 2005. 

George Kontopidis: If I was you the first thing I will try is to increase protein concentration. Also after crystal formed you can take the solution to room temperature that also may help.With the existing crystals just open the container or the vial where the crystals formed to allow further evaporation ( do not let it dry out) which will result to increase of protein concentration and hopefully increase crystal size. 

A.ErtugrulCansizoglu: You might try vapor diffusion against water. As such you keep you protein sample in a mild-salty condition- where it doesn't actually crystallize, then set up your drops using pure water. 
This worked for a case in our lab. Slowing down the crystallization made the crystals bigger. 

Sameer Velankar: I once crystallized a protein in similar conditions. For that matter many Thymidylate synthase crystallize by decreasing the salt concentration. You can add very little salt (15-25mM) in the drop and keep water or 2-5mM salt in the well. This controls the growth of the crystals and if you are lucky you will get bigger crystals. 

Christopher F. Snook:I would suggest increasing the drop volume, i.e. going from say 4 to 8 microliters. This has a two-fold effect: an increase in the protein available for crystallisation and a reduction in the equilibration of the drop with the well solution. Another possiblity would be to increase the concentration which may increase the nucletaion at the expense of crystal growth. 

deacona@mail.rockefeller.edu: I have never encontered such a situation, but maybe some microdialysis from your storage buffer against pure water may help somewhat? I believe this is a common strategy for antibody crystallization. As usual, micro/macro-seeding, varying protein concentration etc may also be of use. Just some thoughts...P.S. These may be of some interest J Biol Chem. 1970 May 25;245(10):2763-4 A crystallographic investigation of a human IgG immunoglobulin.Edmundson AB, Wood MK, Schiffer M, Hardman KD, Ainsworth CF, Ely KR. 
J Mol Biol. 1997 Dec 19;274(5):748-56. Use of organic cosmotropic solutes to crystallize flexible proteins: application to T7 RNA polymerase and its complex with the inhibitor T7 lysozyme. Jeruzalmi D, Steitz TA. 

Oleg Tsodikov: I would still spin down the crystals (or dissolve them in the presence of small amount of salt or glycerol in and some buffer) and then screen in a variety of different conditions. 

David Aragao: We had, some years ago, a very similar case (salting inn?). Our crystals started to appear in the purification process everytime the ionic strenght decreased. The final crystals were big and we got 1.2 Å resolution from them. I know there are nice comercial dialysis buttons nowadays but we made our ones at the time. Maybe the description of our optimization process can help you (see materials and methods). Acta Crystallogr D Biol Crystallogr. 2003 Apr;59(Pt 4):644-53. 

jjwarren@duke.edu: I would recommend that you check out Jeruzalmi, D. and Steitz, T.A. (1997) Use of organic cosmotropic solutes to crystallize flexible proteins: application to T7 RNA polymerase and its complex with the inhibitor T7 lysozyme. J. Mol. Biol., 274, 748-56. Their methods are generally applicable, but they focus heavily on proteins that crystallize at low ionic strength... 

Christopher Colbert: Seriously, you might want to consider screening for additives. 

Ewa Skrzypczak-Jankun: Try salt - increasing the ionic strength often makes crystals more bulky. Also adding salt will change saturation point and can slow crystallization If nothing else NaCl is CHEAP. Do you have any information about affinity to ions? what your protein likes and dislikes? digging in the literature can provide valuable clues. 

Stephen Prince: You could t

a. The protein may have a high degree of structural flexibility and therefore structural heterogeneity. Try screening ligands, additive, co-factors and other small and large molecules that might help the protein assume a more rigid conformation. 

b. Structural variations in the protein due to enzymatic modification, partial deamination, partial oxidation, other post translational modifications can lead to sample heterogeneity. Analyze your sample and conditions from expression to solubilization to purification to crystallization to identify what might lead to sample heterogeneity. Remove it or change it. If working with an enzyme, bind an inhibitor or ligand to the enzyme in an attempt to lock into a more rigid and homogeneous conformation. Reduce the number of steps and the amount of time between expression and crystallization. 

c. If the crystal is growing overnight, the high rate of crystal growth may indicate a high level in the number of defects in the crystal lattice. Slow the rate of crystal growth. Reduce reagent concentration or protein concentration. Layer silicon oil over the reagent reservoir to slow vapor diffusion. Use less reagent in the reservoir. Evaluate drop dilution and drop ratios. For example, dilute the protein 1:1 with your sample buffer. Set vapor diffusion experiments with diluted sample:reagent drop ratios of 1:1, 2:1 and 3:1. The final protein concentration will remain similar between the original and diluted samples but the path of equilibration with be different and the rate will be lower than for the original drop. 

d. Mount the crystal, pre cryo, in a capillary and test for diffraction to confirm that cryo is not causing the poor diffraction. If room temperature data is better than cryo, your cryo method and cryo procedure might benefit from further refinement. 

e. Screen additives. 

f. Screen crystallization kits as additives. Mix 70 to 85 microliters of the optimized crystallization reagent with 30 to 15 microliters of solutions from your favorite Hampton Research screen (Crystal Screen, Crystal Screen 2, Index, SaltRx, etc.) to create a 100 microliter solution. Mix thoroughly. Pipet 100 ul of this solution into a 96 well sitting drop plate. Create a drop mixing equal amounts of reagent and sample, seal and allow to equilibrate. Varying the ratio of optimized reagent: screen reagent and drop ratio can also be tried. Watch out for false positive salt crystals as these random mixes may create insoluble salts. Look in the reservoir for salt precipitates or crystals. If one prefers to use 24 well plates, simply change the ratio of optimized crystallization reagent:screen reagent. For example, in a Cryschem plate try 350-450:150-50. In a VDX Plate try 750-900:250:100. 

References: 
1. Personal communication with Annie Hassell at GlaxoSmithKline, circa 1996. 

2. Acta Cryst. (2005). D61, 646–650. 
Crystallization of foot-and-mouth disease virus 3C protease: surface mutagenesis and a novel 
crystal-optimization strategy. James R. Birtley and Stephen Curry.

Desiccating, or drying a protein crystal can be a way to salvage or improve the diffraction quality. The procedure consists of removing a non-diffracting crystal from the X-ray beam, plunging it into a soaking solution made of the original crystallization reagent supplemented with a suitable cryoprotectant (glycerol, ethylene glycol, MPD, PEG 400, etc.) and then allowing the drop to dry in the evaporating sitting or hanging drop for 15 minutes to several hours. Try 1 ul cryo reagent plus 9 ul of original crystallization reagent for the drop. Remount the treated crystals and examine for diffraction. The procedure is originally described by Chantal Abergel. Reference: Spectacular improvement of X-ray diffraction through fast desiccation of protein crystals. Chantal Abergel. Acta Cryst. (2004). D60, 1413-1416.

Impurities adsorbing onto the crystalline surface can prevent further growth of a crystal. Repair and growth of an impurity poisoned crystal is possible by using a dissolve and restore protocol described by Plomp et al (2003). By temporarily applying undersaturation conditions, an impurity adsorbed layer can be removed, which followed by saturation conditions can revive crystal growth with the potential for crystal growwth without the incorporation of impurities. Regular cycles of short periods of undersaturation followed by longer periods of supersaturation can be applied to dissolve and restore, with the potential of a larger crystal with fewer impurities and defects. Temperature manipulation (cycling) for samples with temeprature dependent solubility is a convenient approach. Altering and cycling sample and reagent concentration is another approach (Heinreichs et al (1992), Koeppe et al (1975), Przybylska (1989)). 

References: 
Plomp et al (2003). Repair of impurity-poisoned protein crystal surfaces. Proteins: Structure, Function, and Genetics 50:486-495. 

Heinreichs et al (1992). Growth of single protein crystalsin a periodically solubility gradient: description of the method and first results. J. Cryst. Growth 122:186-193. 

Koeppe et al (1975). A pulsed diffusion technique for the growth of protein crystals for X-ray diffraction. J. Mol. Biol. 98: 155-160. 

Przybylska (1989). A double cell for controlling nucleation and growth of protein crystals. J. Appl. Cryst. 1989:22:115-118. 

Heavy atom derivatives can some times be difficult to obtain in the presence of sulfate salt crystallization reagents such as ammonium sulfate or lithium sulfate. 

However, difficult does not mean impossible, so be sure to screen heavy atoms in the salt based condition before trying to cure a problem that might not exist. 

When one is unable to obtain succcessful derivatives in the presence of sulfate salt based crystallization reagents, one might try transferring the crystals to high concentrations of a similar acetate salt based crystallization. For example, ammonium acetate instead of ammonium sulfate, or lithium acetate instead of lithium sulfate. This will maintain the high ionic strength of the crystallization reagent yet increase the solubility of some heavy atoms. Acetate, in higher concentrations can also act as a cryo salt. 

Try replacing the original salt with sodium malonate or Tacsimate. Malonate: a versatile cryoprotectant and stabilizing solution for salt-grown macromolecular crystals. T. Holyoak, T. D. Fenn, M. A. Wilson, A. G. Moulin, D. Ringe and G. A. Petsko. Acta Cryst. (2003). D59, 2356-2358 

Try halide soaking using KI or KBr. Novel approach to phasing proteins: derivatization by short cryo-soaking with halides Z. Dauter, M. Dauter and K. R. Rajashankar Acta Cryst. (2000). D56, 232-237. 

One might also consider trying 100 mM cobalt hexammine or iridium hexammine as derivatives in the presence of acetate salts. 

Try crosslinking. A gentle vapor-diffusion technique for cross-linking of protein crystals for cryocrystallography. Carol J. Lusty. J. Appl. Cryst. (1999). 32, 106-112.

Problem: Enzyme crystals grown in ammonium sulfate having trouble binding ligand. 

Solution: This is not an unusual situation with sulfate, and, yes, sulfate often occupies phosphate binding sites, particularly when you're work with 1-3 molar concentration of ammonium sulfate. The simplest way to remove sulfate from the crystal is to transfer to high concentrations of citrate. From my own graduate work on GPDH in the 1970's, we just transferred the crystals from 3.0 M AS to 1.44 M Na citrate. While the crystals weren't stable in citrate for long periods of time (days), that is not an issue today with cryocrystallography and flash-freezing. You might even try formate or malonate, which can have a modest cyroprotectant behavior. Another, and perhaps more relevant, method is the one devised by Bill Ray in the late 1980's for phosphoglucomutase (PGM). Apo-PGM was crystallized in high AS, which caused the same problems as you are experiencing. Ever the perfectionist, Bill devised a systematic method to transfer PGM crystals from AS to a PEG solution that allowed the formation of enzyme complexes (Biochemistry 30, 6866, 1991). Bill also looked at glycine as a replacement as well. So there are a number of ways to "desalt" crystals without resorting to crosslinking, while preserving good diffraction characteristics. Michael Garavito, Michigan State University .

Using the crystallization reagent that produced the initial hit, or reagent formulations which have been partially optimized, set additional crystallization experiments using other methods and techniques. If sitting drop vapor diffusion was used to produce the initial hit, try hanging drop vapor diffusion, microbatch, free interface diffusion, or dialysis. Try different plates and different apparatus using the same method to discover whether the difference in equilibration will produce a better crystal, a crystal in a shorter period of time, or a different crystal form. The volume of the chamber or well, the space between the drop and the reagent, the shape of the chamber and the well and also the method and time required to set the experiment may affect the outcome of the crystallization experiment. 

The path is as important as the start point and end point in determining ideal nucleation and growth conditions.

If the crystallization reagent producing a hit is a complex mixture containing, for example, salt, detergent, additive, metal ions, ligands, reducing agent, EDTA, cryogen, or other chemical, prepare optimization reagents based on the hit, but systematically omit each of the components one at a time. Which chemicals appear to make a difference in the crystal size, number and quality? Omit from future optimization experiments any chemical which appears to be irrelevant. Identify and remove irrelevant chemicals before adding the evaluating new chemicals. As with the sample buffer, keep the crystallization reagent as simple as possible. Include only chemicals which promote and enhance sample stability, homogeneity and crystallization.

The skill of crystal optimization of a crystal is an art form that improves with effort, thought, discussion, reflection, reading, learning, trial and error, patience, tinkering and time. 

The path of optimization is rarely linear and the variables affecting crystallization are not independent. This makes it impossible to sample all crystallization variables and evaluate them at fine intervals. 

Optimization may follow a main trail, but there are typically many branches in the trail to consider, explore and evaluate. And typically time and sample availability do not allow one to explore each and every branch. 

So one is left with the question of what they should try or what they should do next to increase the size or improve the quality of of the crystal. And there are usually plenty of recommendations to consider. But which recommendations should be considered and tried and in what order or priority, and which ones should be ignored for now and tried later, or perhaps discarded altogether? 

It is the art of experienced crystal grower to consider the results of systematic experiments of many different samples over much time and use this knowledge based to synthesize the best plan for growing perfect crystals. 

Optimization can be tedious and require numerous iterations before one uncovers which combination chemical and physical variables produce the perfect crystal. 

Optimization is an art and a technical skill. As such it is dynamic and improves in direct proportion to the number of challenges, the degree of the challenges and experience. 

A good crystal grower never is, they simply get better and better and better... 

A skilled craftsman knows they cannot do their job without the proper tools. Maybe some jobs can be accomplished with one or two common and simple tools, but a skilled craftsman, over time, will build not only experience but also a tool box. A tool box of both general and specialized tools that will allow them to handle all tasks in an efficient manner. And over time a skilled craftsman gains appreciation of the importance that quality, reliability and support bring when deciding which tools to acquire. 

A crystal grower also needs to build a tool box. This tool box will consist of apparatus and devices, plates and seals, kits and reagents; some general, some specialized. Initially the crystal grower will want o acquire tools that have general utility. Later, more specialized tools can be added so that when the need arises, these tools will be available to use and evaluate. The access to a useful and broad portfolio of tools ensures the crystal grower will have access to evaluate those variables which could lead to the perfect crystal. 

You can only use and evaluate what you have and know how to use. 

Phase separation in a crystallization experiment 

What are these droplets in my crystallization experiment? 
Sometimes when a crystallization reagent is added to a protein the result is liquid drops. This liquid-liquid phase separation is also referred to as “oils” or “phase separation” (phase sep). Phase separation can be observed by simply changing pH, temperature or reagent conditions and concentration. Phase separation droplets typically contain a high concentration of protein. Eventually, with time or centrifugation, the drops may separate completely from the rest of the solution and form two liquid phases. The concentration of the various components will be different in each of the two phases. 

Phase separation droplets may be numerous or small or few and large, depending upon solution conditions and time. Phase separation may exist for days, weeks or months. 

What to do? 
The appearance of phase separation in a crystallization experiment can be an indication of a metastable solution, existing somewhere between the stable undersaturated (clear) and the unstable precipitation zone. Phase separation is a metastable transition, a position in the protein crystallization phase diagram crystals may form. Physical, chemical and biochemical variables should be evaluated for their ability to create an activation and event to nucleat crystal growth in phase separation. 

Approaches to promote crystallization from phase separation: 

Alter temperature 
Evaluate temperatures between 4 and 37 degrees Celsius increment of 5 degrees. 

Alter reagent concentration 
Increase and decrease concentration 10-20% in 2% increments. 

Alter pH 
Increase and decrease 1-2 pH units in 0.2 increments. 

Screen small molecules to perturb sample-sample and sample-solvent interactions 
Additive Screen 
Detergent Screen 
Silver Bullet Screen 

Alter protein concentration and/or change drop ratio 

Proteolytically modify the sample.

Avoid buffers that bind metal ions such as zinc. TRIS and MES have been used successfully as buffers for the crystallization of zinc finger proteins. 

If a histidine is present in your zinc finger protein, do not go below pH 5 or you will lose the zinc binding from the protein. 

High pH, up to 10 is okay, but keep the concentration of reducing agent at or below 1 mM. 

Do not use EDTA or other metal ion chelators in the prepapration, purification or crystallization. 

Reference: Ray Brown, 3DBioScience, ccp4bb January 28, 2011.

Native PAGE, mass spectrometry and analysis of X-ray data. 

Screening for phasing atoms in protein crystallography. Boggon TJ, Shapiro L. Structure. 2000 Jul 15;8(7):R143-9. 

Mass-spectrometry assisted heavy-atom derivative screening of human Fc gamma RIII crystals.Sun PD, Hammer CH. Acta Cryst. (2000). D56, 161-168 doi:10.1107/S0907444999015188.

Buffer Type & pH Case Study Using the Slice pH screen 

• Lipase B, 20 mg/ml in deionized water. Drop: 0.5 ml Lipase B + 0.5 ml Slice pH / Reservoir: 50 ml 3.0 M Sodium chloride 

• Sitting Drop Vapor Diffusion MRC 2 Well Crystallization Plate (Swissci) HR3-083 

Results: 

• Three different results using three different buffers at the same pH (see images above). 
pH 4.5 DL-Malic acid Crystals; pH 4.5 Sodium acetate Clear; pH 4.5 Sodium potassium phosphate Precipitate 

• Glycine pH 8.6 Clear; Glycine pH 8.8-9.5 Precipitate/Phase Separation; pH 9.6 Crystals/Precipitate 

Recommendation: 
Crystallization screen should focus on DL-Malic acid as a buffer / precipitant near pH 4.5 as well as Glycine as a buffer/precipitant between pH 8.6-9.6. Previously reported in the literature: Lipase is most stable between pH 5 to 7 with an isoelectric point of 6.0. Typical pH range for crystallization is 4 to 6. (The sequence, crystal structure determination and refinement of two crystal forms of lipase B from Candida antarctica. Uppenberg, J., Hansen, M.T., Patkar, S., Jones, T.A. Structure v2 pp. 293-308, 1994; Crystallization and preliminary X-ray studies of lipase B from Candida antarctica. Uppenberg J., Patkar S.; Bergfors T., Jones, T. A.. Journal of Molecular Biology, 1994, vol. 235, no2, pp. 790-792).

Try one or more of the following to slow down the rate of crystal growth when trying to improve crystal quality. 

Increase drop size. Larger drops equilibrate more slowly than smaller drops. 

Add a small amount of glycerol (1-5% v/v) or other reagent that might solubilize the sample. Reagents to consider include chaotropes, kosmotropes, detergents, and salts. 

Cover the reservoir solution with paraffin or silicon oil or a mixture of both in order to reduce the rate of vapor diffusion between the drop and the reservoir. 

Dilute the drop. This can suppress excessive nucleation for and increase crystal size. For example, instead of setting drops with 1 uL of protein and 1 uL of reservoir, try setting the drop with 1 uL of protein, 1 uL of water and 1 uL of reservoir. One can increase the water volume even further, as needed to control nucleation and the rate of crystal growth.

When experiencing difficulty obtaining high quality SeMet crystals, try using the native crystals for streak seeding and the SeMet protein for the drop. 

Reference: Structure of Escherichia coli BamD and its functional implications in outer membrane protein assembly. Cheng Dong, Hai-Feng Hou, Xue Yang, Yue-Quan Shenb and Yu-Hui Dong. Acta Cryst. (2012). D68, 95–101.

Crystals growing in 1-propanol could be screened against the following reagents. 

1-butanol 
Ethanol 

Maybe you're trying to get away from the fun of mounting crystals in volatile reagents? In that case you could try MPD and glycerol or other polyols although these reagents can be a bit different in action than the volatile alcohols. But one never knows until it's tried. 

If your reagent is only 1-propanol and buffer and no other salts or additive, this would be a very low ionic strength reagent. In such instances one should consider screening temperature and pH since these two variables can be more significant in low ionic strength environments. 

And perhaps the following as additives if the 1-propanol you're using it also mixed with a salt to create the crystallization reagent. Try different chain length alkanediols to see how the change in alkanediol affects sample-sample and sample-solvent interactions as well as sample solubility. 

Butane-1,4-diol 
butane-1,2-diol 
pentane-1,2-diol 
pentane-1,5-diol 
hexane-1,2-diol 
hexane-1,6-diol 
heptane-1,7-diol 
octane-1,2-diol 

Reference: Roles of Additives and Precipitants in Crystallization of Calcium- and Integrin-Binding Protein. Berger et al. CRYSTAL GROWTH & DESIGN 2005 VOL.5,NO.4 1499-1507 

One might also try rescreening with their preferred crystallization screen but adding 10% 1-propanol into the screen reagent. Meaning, make up your reservoir solution with 90% screen reagent and 10% of 100% 1-propanol. For example, a 100 ul reservoir would contain 90 ul of screen reagent and 10% of 1-propanol.

The technical mumbo jumbo first. The physical properties of isotropic materials, such as glasses, liquids and amorphous materials, do not depend on direction. However, most properties of a wide variety of crystals (including liquid crystals) do show such variation. This anisotropy of physical properties originates in the anisotropic build-up of the materials (crystal structure). Anisotropy in the optical properties of uniaxial crystals is referred to as either birefringence or dichroism, depending on whether the index of refraction or the absorption coefficient is concerned. Birefringence means that there are two distinct speeds with which light can propagate, depending on the direction of propagation. When a light ray splits into two beams as it passes through a material, the effect is called birefringence (or double refraction) and the material is birefringent. If you look at something through a birefringent material, you'll see double. The word birefringence comes from the Latin bi- (twice) plus refringere (to break up). Thus, the light rays are "broken in two" by a birefringent material. One well-known example of a birefringent medium is crystalline calcite (calcium carbonate). If you look at the world through a clear crystal of calcite (calcium carbonate), you will see double. Place such a crystal on a drawing, and you'll see two overlapping copies of the drawing. The molecular structure of calcite causes double refraction, in which each light ray is split into two rays that emerge from the crystal at slightly different angles. Calcite shows this more clearly than most crystals, but quartz and many other crystalline minerals also split light ray. 

Now, the practical interpretation for crystal growers. You might hear the word birefringence used quite often by crystal growers when viewing crystals under a microscope. Here, crystal growers are stretching the definition of the term birefringence to describe the colorful display produced by biological macromolecular crystals when polarized light is passed through the crystal. 

A light microscope with polarizing optics is required to observe birefringence. The following path is a typical set-up. Light passes from the light source through the first polarizing lens, then the specimen (crystal) then the second polarizing optic, the magnifying optics and then into your eye. On many typical polarization set ups, the second polarizing filter can be rotated while the specimen is stationary. Rotating the polarizing optic without something to rotate the plane of polarized light in the path (i.e. a crystal) will result in one seeing light, dark, light, dark, as the filter is rotated. But if a crystal with birefringent properties (i.e. a biological macromolecular crystal) is positioned in between the two polarizing filters, one will observe changing colors as the polarizing filter is rotated. Specifically, when the polarizing filters are aligned such that the field is dark, a birefringent object (crystal) will glow with color. 

Birefringence is one way we can differentiate amorphous precipitate from microcrystals in a drop when viewed under a microscope. Precipitate does not have birefringent properties while most biological macromolecular crystal do have birefringent properties. 

One drawback with using birefringence in today's crystal growth world is that most of the crystallization devices utilized are made from plastic such as polystyrene and polypropylene. These plastics are optically active can be birefringent. In fact, often times the colors we see displayed in crystals are contributions from the plastic birefringence. However, it is still possible to observe microcrystalline birefringence in the plastic trays, but there is usually a contributory effect from the plates used to grow the crystals. One way to avoid this is to grow crystals in a glass device or at least observe the crystals in a path that is free of plastic. 

Birefringent precipitates will glow, sparkle, or glisten. 

To test for birefringence, position the polarizers so the field of view is dark WITHOUT they crystallization plate or set up. Place the tray into position on the microscope. If a crystal is birefringent, some of the light passing through the crystal will be rotated and passed through the second (analyzing) polarizing filter. The intensity of the transmitted light will increase and decrease as the crystal is rotated or the polarizer is rotated. Remember, birefringence is not ALWAYS clearly visible when plastic is in the light path (i.e. when you use plastic slides or crystallization plates). However, a birefringent crystal viewed in a plastic tray or plastic cover slide will have a different color than the background (i.e. plastic plate) or foreground (plastic slide). Finally, birefringence is a property of crystals, both biological (proteins, peptides, and nucleic acids) and inorganic crystals (salts). Birefringence is MORE pronounced in inorganic (salt). 

A quick comment on what to do with birefringent precipitates. Streak seeding is a common and often successful method of taking advantage of microcrystalline precipitate to grow large single crystals. But that starts with an S so we cannot talk about that this time!

Chaotropic (order-breakers) substances tend to increase the solubility of macromolecules in water. Chaotropic substances decrease the ordered structure of water. Chaotropes lower the cohesion between water molecules. Chaotropes lower the surface tension of water. In general, the surface tension decreases as the chaotropy increases. Chaotropic salts can perturb sample sample and sample solvent interactions by shielding charges and preventing the stabilization of salt bridges. Hydrogen bonding is stronger in nonpolar media, so salts, which increase the chemical polarity of the solvent, can also destabilize hydrogen bonding because there are insufficient water molecules to effectively solvate the ions. This can result in ion-dipole interactions between the salts and hydrogen bonding species which are more favorable than normal hydrogen bonds. Chaotropes break down the hydrogen-bonded network of water, so allowing macromolecules more structural freedom and encouraging protein extension and denaturation. However, lower, non-denaturing concentrations of chaotropes can be used to manipulate sample sample and sample solvent interactions as a means to promote crystallization or improve crystals. 

Chaotropes include urea, guanidine, tetramethylammonium chloride. guanidine thiocyanate, thiourea, lithium perchlorate, and sodium thiocyanate.

Vary the sample:reagent drop ratio. A typical sample:drop ratio is 1:1. Try a sample:drop ratio of 1 part sample:2 parts reagent as well as 2 parts reagent:1 part sample. 

Varying the drop ratio will change the initial and final sample concentration and the initial reagent concentration and also alter the equilibration kinetics between the drop and the reservoir. 

Varying the drop ratio can be helpful when trying to reproduce conditions when changing crystallization methods and can also be a useful screening and optimization variable.

Ethylene glycol. Also known as 1,2-Ethanediol as well as glycol. Molecular mass of 62.07. HOCH2CH2OH. A nice little polyol that can be useful as a cryoprotectant. Typically used as a cryoprotectant in the concentration range of 10 to 30% v/v. Also useful as a non volatile organic additive when used in the concentration range of 1 to 5% v/v and in some cases, much higher (15 to 25% v/v). As an additive, ethylene glycol has been reported to sometimes increase the size and quality of the crystal. Finally, it is useful as a crystallization reagent in the concentration range of 25 to 30% v/v. It is even thought that ethylene glycol can have a structure-stabilizing effect and may be useful in the crystallization of flexible proteins.

Fab and Fv fragments can be used as co-crystallization agents. Fab fragments have decent solubility properties and bind specifically to antigens with reasonable equilibrium constants (105 to 108 M-1). Fab fragments can, in some instances, effectively transform aggregated protein into soluble, monodisperse protein, suitable for crystallization trials. Since antibodies are frequently available from related biochemical studies, they can often be applied more readily than molecular biology (point mutants, truncations, molecular engineering). However, it seems MOBO gets faster and easier with each passing year. It is often useful to consider screening several different Fab fragments that recognize different epitopes for crystallization trials. Sometimes Fab fragments can immobilize a region of a protein sample, reducing sample flexibility, enhancing conformational homogeneity of the sample which in turn can enhance chances for crystallization. 
Using an Fv fragment in lieu of a Fab fragment might have some advantage since there is no flexible elbow to inhibit crystallization. 
When preparing antigen-antibody complexes for crystallization, one should carefully select the antibody, prepare a homogeneous Fab species, and prepare the Fab-sample complex with proper stoichiometry. Ideally the Fab should not interfere with the native sample conformation or activity. 

The presence of a Fab fragment in the crystal can assist in the crystallographic structure determination by allowing one to utilize molecular replacement. 

Fab and Fv fragments should be considered useful tools in the crystallization toolbox for manipulating sample solubility, conformational flexibility, and crystal lattice contacts, as well as being a possible tool for molecular replacement. 

Reference: 
The use of antibody fragments for crystallization and structure determinations. LC Kovari, C. Momany and MG Rossman. Structure 15 December 1995, 3: 1291-1293.

1,6 Hexanediol (C6H14O2 Mr 118.18) is a non-volatile alcohol which most likely precipitates biological macromolecules by lowering the chemical activity of water. Non-volatile organics can be effective crystallization reagents for proteins and are particularly appropriate for nucleic acids. This non-volatile alcohol steals water molecules from the biological macromolecule through hydrogen bonding and reduces the electrostatic screening effectiveness of the water. Typical stock solutions of 6.0 M (about 71% w/v) are typical for this crystallization reagent. Typical reservoir concentrations for this reagent are 1 to 4 M across a broad pH range. 1,6 Hexanediol can be utilized in the presence of numerous anions and cations, as well as a broad range of salts. It can be a substitute crystallization reagent for MPD, PEG 400, and PEG MME 550. It can also be utilized as an additive in the concentration range of 0.1 to 0.2 M. 1,6 Hexanediol is a waxy solid or flake. When formulated with water, the reaction is endothermic and one might need to warm the solution to effect complete dissolution at high levels of supersaturation. 

1,6 Hexanediol is utilized as a precipitant in Crystal Screen 2 and is available as a preformulated, sterile filtered reagent from Hampton Research (Catalog Number HR2-625; 6.0 M 1,6 Hexanediol, 200 ml).

Nucleases can be a real pain during the crystallization of nucleic acids. Here are some nasty things nuclease can do to your sample: Modification of sample size, modification of the charge or hydrophobicity, partial or total loss of activity, or utter desctruction of the sample. Traces of nuclease can be difficult to detect even when overloading electrophoresis gels and can obviously cause sample damage during purification, concentration, and storage of samples. What is one to do? Well, one can include a nuclease inhibitor in the prep or sample to protect the sample from ribonuclease and deoxyribonucleases. Inhibitors of nucleases include RNasin (from Promega), ribonucleaoside-vanadyl complexes, and DEPC. Inhibitors of deoxyribonucleases include DEPC and chelators such as EDTA or EGTA.

The most popular volatile organic solvents used in biological macromolecular crystallization have been ethanol, acetone, isopropanol, tert-butanol, 1,3-propanediol, acetonitrile, DMSO, methanol, and 1,3-butyrolactone. Organic solvents can be utilized as a primary precipitant (buffered or unbuffered), as a secondary precipitant in the presence of salt or polymer (primary precipitant), or as additives. The most popular nonvolatile organics have been MPD and 1,6-hexanediol. Organic solvents act as precipitants by lowering the chemical activity of water. This means they steal water molecules from biological macromolecules in solution, through a process of hydrogen bonding. This in turn reduces the dieletric constant of the solution. Current popular thought is that organic solvents should be used at low temperatures (4°C or lower) and at the lowest possible ionic strength, keeping in mind to include whatever is necessary to stabilize the sample (buffer, divalent cations, etc.).

pH and Protein Crystallization 

While biological macromolecules are large and provide the potential for many interactions to occur, a remarkable feature of protein, nucleic acid and virus crystals is the relatively few intermolecular interactions that are actually observed in crystals whose structures have been solved. 

Salt bridges, electrostatic interactions, hydrogen bonds, hydrophobic interactions, Coulombic interactions, and Van der Waals forces are examples of the few intermolecular interactions involved between molecules forming the crystal lattice. 

The alteration and variation of dielectric environment of the crystallization reagent can be an important variable in the initial attraction, orientation and attachment of molecules forming a crystal. It is partly for this reason that type and concentration of salt and precipitant, pH, and additives are varied in crystallization screens. So it is logical to pursue the alteration and refinement of these variables in the optimization of crystal size and quality. 

Salt bridges between protein molecules require protonation of the basic residues and deprotonation of the acidic groups in order to form. Neutralization of either, as happens at low and high pH, disrupts those interactions, or at least lessens their rate and frequency of formation. Thus, the manipulation and control of the relative number of charged groups and their capacity to engage in electrostatic interactions may be used to control the size and quality of the crystal. To accomplish this, we vary the pH of the crystallization reagent. 

pH is an effective crystallization variable because most proteins demonstrate pH dependent solubility minima and will solubilize, precipitate, or crystallize at particular pH values. The solubility minima may correspond with the isoelectric point (pI) of the protein, but this is not always the case. The solubility minima is often complex and may depend on other chemical and physical variables in the crystallization experiment. 

A near neutral pH 6.5-7.5 is the most frequently reported pH value for protein crystallization. A plot of reported crystallization pH between 2 and 10 is nearly bell shaped with the peak around 7. So is entirely reasonable to perform initial screens covering pH 4 to 9 as this is the reported range of pH used to crystallize most proteins. Since protein crystallization has been reported as low as pH 2 and as high as pH 10 one should consider screening these pH extremes, as appropriate, during optimization or when preliminary screens produce anything but crystals. 

The impact of pH is magnified as the ionic strength is reduced. 

Crystal morphology can be highly pH dependent1. 

How much and how little should one vary the pH during optimization? Increase and decrease the pH of the crystallization reagent until no crystals are observed. Then perform a fine pH screen between the pH of the screen reagent producing the crystals and the pH where crystals no longer form2. Remember, the difference between clear drops, amorphous precipitate, microcrystals and larger single crystals may only be a few tenths of a pH unit apart. For example, if your crystallization reagent is pH 6.5 your optimization reagents might screen plus or minus 1 to 2 pH units above and below pH 6.5 in 0.5 pH increments (5.5-7.5 or 4.5-8.5). Let’s say you observe crystals at 6.5, 7.0 and 7.5 but not at 8.0 and 8.5. You might then screen between pH 7.5 and 8.0 or even 8.5 in 0.1 pH increments to precisely control the relative numbers of charged groups and their capacity to engage in electrostatic interactions.

Should I use the same buffer as the screen buffer used to produce the crystal? Yes and no. Yes, be sure to screen pH using the same buffer as that used in the original screen reagent. But also consider screening other buffer salts, appropriate for the pH range screened. For example, if your screen reagent buffer is HEPES pH 7.0, include HEPES in your optimization and also consider buffers such as ACES, ADA, Bis-Tris, Imidazole, PIPES and other buffers with a pK covering pH 7.0. 

What concentration of buffer should be used for crystallization reagents? A 0.1 M buffer concentration in a crystallization reagent is frequently used. A 0.1 M final buffer concentration is typically of sufficient concentration to manipulate and control the pH of the experiment and allows the use and dilution of a 1.0 M buffer stock during reagent formulation with plenty of volume remaining for the addition of other chemicals (salts, polymers, additives, etc) to the reagent. When truly low ionic strength crystallization reagents are desired, the buffer concentration in the reagent can be dropped to 0.05 to 0.01 M. But be consider the pH and buffer concentration in the sample and keep the reagent buffer at or above that of the concentration of the sample buffer so the reagent can drive and control the pH of the experiment and not the sample buffer. 

Are we really controlling pH or just manipulating pH? I have a 1.0 M buffer stock adjusted to pH 7 and when it was diluted to 0.1 M and added to PEG and salt the pH ended up being 6.5. Am I fighting a losing battle? Yes, we are controlling and manipulating pH but not fighting a losing battle. Salts and additives can have strong buffering properties and push the actual measured pH of the reagent far from that of the original buffer pH. This affect will vary with the type of salt or additive, the concentration of the salt or additive, the buffer and the pH of each chemical in the reagent. Unless a pH titration is performed after all of the chemicals are added to the reagent, it is likely the final measured pH will be different than that of the sample buffer. Some times these pH changes are subtle and some times they are very significant. 

Some reagent solutions, such as those in the Hampton Research Grid Screen kits do have their pH titration performed following chemical addition. This is done so the experimenter can test exact pH values under a variety of precipitants at different concentrations. However, most screen reagents begin with a titrated buffer stock that is then diluted with other chemicals and water with no final pH adjustment. One strong argument for using the latter formulation is the ease, convenience and speed of formulation, ease of reproducibility and ability to use automation without involving pH titration. 

So while perhaps one does not worry obsessively about anticipated versus actual pH, one must be extremely aware of and concerned with the fastidious formulation of buffers using high purity chemicals, quantitative methods, properly maintained and calibrated instrumentation. This will help to ensure control and reproducibility of the crystallization of the sample. 

What buffer concentration and pH should I use for my sample? Use a buffer concentration between 10 and 20 mM so that the crystallization reagent, typically at 100 mM, can manipulate and control the pH of the experiment. If the sample is not soluble in low ionic strength, add 25 to 200 mM sodium chloride. One might also consider what ligands, metals. co-factors, reducing agents, chelating agents or other additive might be useful in promoting, enhancing and stabilizing sample stability, homogeneity and monidispersity. pH is often the single most important parameter for sample solubility. One should screen a range of pH between 2 and 10 to determine the ideal pH for sample solubility. If you do not have time to screen pH, know nothing about the sample and are forced to choose a buffer, choose HEPES sodium pH 6.8. 

What buffers should I avoid and why? Inorganic buffers can work wonderfully but can also produce salt crystals, especially at low temperature, in the presence of metals and salts and high concentrations of precipitants. Citric acid is a chelator of many metal ions and might be a poor choice when screening metal additives and heavy metals. Cacodylate is an arsenic compound, is poisonous and should be handled appropriately. 

References 

1. Structural studies on the adenovirus hexon. Franklin RM, Harrison SC, Pettersson U, Philipson L, Brändén CI, Werner PE. Cold Spring Harb Symp Quant Biol. 1972;36:503-10. 

2. Increasing the size of microcrystals by fine sampling of pH limits. A. McPherson. J. Appl. Cryst. (1995). 28, 362-365. 

Acid pH crystallization of the basic protein lysin from the spermatozoa of red abalone (Haliotis rufescens). T. C. Diller, A. Shaw, E. A. Stura, V. D. Vacquier and C. D. Stout. Acta Cryst. (1994). D50, 620-626. 

Protein Isoelectric Point as a Predictor for Increased Crystallization Screening Efficiency. K. A. Kantardjieff and B. Rupp. Bioinformatics 20(14): 2162-2168 (2004). 

Distributions of pI vs pH provide strong prior information for the design of crystallization screening experiments. K. A. Kantardjieff, M. Jamshidian and B. Rupp. Bioinformatics 20(14): 2171-2174 (2004). 

Novel buffer systems for macromolecular crystallization. J. Newman. Acta Cryst. (2004). D60, 610-612. 

Optimization of buffer solutions for protein crystallization. R. A. Gosavi, T. C. Mueser and C. A. Schall. Acta Cryst. (2008). D64, 506-514. 

Optimum solubility (OS) screening: an efficient method to optimize buffer conditions for homogeneity and crystallization of proteins. J. Jancarik, R. Pufan, C. Hong, S.-H. Kim and R. Kim. Acta Crystallographica Section D, Biological Crystallography, Volume 60, Part 9 (September 2004). 

Crystallization Optimum Solubility Screening: using crystallization results to identify the optimal buffer for protein crystal formation. Bernard Collins, Raymond C. Stevens, and Rebecca Page. Acta Crystallogr Sect F Struct Biol Cryst Commun. 2005 December 1; 61(Pt 12): 1035–1038. 

Introduction of Fluorometry to the Screening of Protein Crystallization Buffers. Takamitsu Ikkai and Katsuhiko Shimada. Journal of Fluorescence, Volume 12, Number 2, June, 2002, Pages 167-171. 

Lepre, C. A., Moore, J. M.; Microdrop screening: A rapid method to optimize solvent conditions for NMR spectroscopy of proteins; Journal of Biomolecular NMR, 12: 493-499, 1998. 

Optimization of Met8p crystals through protein-storage buffer manipulation. H. L. Schubert, E. Raux, M. J. Warren and K. S. Wilson. Acta Cryst. (2001). D57, 867-869. 

The Effect of Temperature and Solution pH on the Nucleation of Tetragonal Lysozyme Crystals. Russell A. Judge, Randolph S. Jacobs, Tyralynn Frazier, Edward H. Snell, and Marc L. Pusey.

Proteolytic modification of proteins can be a tool for making small active fragments of proteins, which might have enhanced solubility characteristics compared to the native protein, which in turn might make the protein more amenable to crystallization. 

However, proteases can also be trouble makers when they are uninvited guests to a protein sample. When protease contamination can be a problem, one can consider adding a protease inhibitor to prevent cleavage of the sample by these nasty proteases. Protease contamination can occur when these nasty beasts are carried over from isolation and purification of the sample from a natural source or expressed proteins. Also, if your lab is situated in an area (on the same floor, building, etc.) where microbial research, plant research, or other work involving the possible generation of fungal and bacterial organisms is possible, then one might consider adding protease inhibitors to the sample for protection. Why? The presence of fungal or microbial contamination in your sample, reagents, or crystallization related plates, capillaries, etc. can lead to growth of such organisms with subsequent release of proteases from these organisms so they can use your innocent little sample for food. Some crystal growers like to include sodium azide or thymol in all their reagents and sample as a deterrent to microbial growth, which prevents the possibility of microbial agents growing and secreting proteases into the sample solution. But sodium azide and thymol can sometimes bind the sample, are toxic, and in some cases do not live well with heavy atoms so some feel it is not wise to simply include these agents as a multi-vitamin in the crystallization cocktail. Rather, clean workspace, sterile filtered samples and reagents, sterile pipet tips, and good technique go a long way to preventing microbial contamination. Well, back to the P for Protease Inhibitors. Here are a few protease inhibitors and their targets. 

Protease: Metalloproteases 
Inhibitor: Chelators like EDTA and EGTA, bestatin, amastatin, thiol 
derivatives, hydroxamic acid, phosphoramidon. 

Protease: Aspartic Acid Proteases 
Inhibitor: Pepstatins and statin derived inhibitors. 

Protease: Cysteine Proteases 
Inhibitor: Thiol binding reagents, peptidyldiazomethanes, epoxysuccinyl 
peptides, cystatins, peptidyl chloromethanes. 

Protease: Most 
Inhibitor: DEPC 

Protease: Serine Proteases 
Inhibitor: Trypsin inhibitors, leupeptin, boronic acids, cyclic peptides, DIFP, PMSF, Pefabloc, aminobenzamidine, 3,4-dichloro isocoumarin, 
chymostatin.

Vibration has been reported to lead to excessive nucleation and crystals of questionable quality. It is often suggested that crystallization experiments be incubated in a location with minimal vibration. Avoid cabinets that are frequently opened and closed, or countertops where equipment such as centrifuges or vortexes live. Incubators can be another sore spot for vibration, especially poorly insulated incubators with compressors that are not well isolated.

Vibration is often associated (in a crystallographers mind) with excessive nucleation. It is popular belief that the best crystals grow when there is NOT a significant source of vibration. It is believed this is particularly important during the optimization and production stages of crystallization. Hence, it is suggested that during optimization and production, one set crystallization experiments and avoid the burning curiosity to look at the plates every single day. Try leaving the setups alone during optimization and production. 
Since vibration can lead to excessive nucleation, it seems one might want to move the plates some during screening. One typically does move plates during screening as it is typically recommended to view experiments each day for the first week and once a week thereafter until the drop dries out. Maybe we need to build vibrators into our screening incubators. Along the same line, it is sometimes observed that plates which have been sitting for some time, then viewed, often have crystals forming in the days following the latent observation. So perhaps one should consider giving the plates a Monty Python nudge-nudge or move them about to stimulate nucleation. 

To avoid vibration, place plates in cupboards, cabinets, or incubators that are infrequently opened and closed. Installing hydraulic door dampers to prevent slamming doors or cupboards is one way to reduce vibration. Some incubators vibrate more than other units and peltier based heating and cooling incubators may be desirable from this perspective. However, some of the circulation fans in these units vibrate quite a bit, so be sure to check out the fan and even consider replacing it with a higher quality unit with less oscillation and vibration.

Reducing agents are substances that cause other chemical species to be reduced or gain electrons. In order for reducing agents to cause the gaining of electrons on some other chemical species, they must undergo oxidation. Therefore reducing agents undergo oxidation when they do their job. 

Dithiothreitol (DTT), beta-mercaptoethanol (beta-me), and Tris(2-Carboxyethyl)-Phosphine Hydrochloride (TCEP HCl) are sulfhydryl protective reducing agents. Reducing agents are typically used to prevent the oxidation of free sulfhydryl residues (cysteines) in the protein. Such oxidation can lead to non-specific aggregation of the sample, sample heterogeneity, inactivity, or denaturation of the sample. In a typical crystallization experiment, reducing agents are used in the concentration range of 1 to 10 mM in the crystallization drop. 

Beta-me has one sulfhydryl group and is the weakest of the three reducing agents discussed here, lasting perhaps two to three days. The supply of beta-me should be replenished every two to three days in the crystallization experiment to maintain the effectiveness of the reducing agent. Since beta-me is volatile, it can be added to the reservoir of vapor diffusion experiments for diffusion into the crystallization drop. 

DTT has two sulfhydryl groups and lasts about three to seven days in a typical crystallization experiment. DTT is not a strong volatile like beta-me (although it does possess a strong odor) and should be added directly to the crystallization drop when possible. Another consideration when working with DTT is to use the dialysis method for crystallization since DTT can be added to the dialysis solution to replenish the supply of reducing agent. 

TCEP HCl is stronger than both beta-me and DTT, lasting about 2 to 3 weeks in a typical crystallization experiment. TCEP hydrochloride can acidify the crystallization solution. Like DTT, TCEP HCl needs to be added directly to the crystallization drop, hence, dialysis is a consideration. 

Beta-me, DTT, and TCEP HCl can behave differently. Like all additives, if you find a class of compounds has an effect on sample stability or crystallization, then screen a variety of compounds in that class to see which one is best for your application. What we are trying to suggest is that like all additives, one might consider evaluating all three reducing agents to see which one works best for your sample. 

There have been reports where reducing agents have been used as successful crystallization additives where there were no free sulfhydryls in the sample. 

Reducing agents can bind metals and trace metal compounds, inactivating the reducing agent and the metal. This can make heavy atom derivatization in the presence of reducing agents, difficult and frustrating. EDTA can be added to the crystallization experiment to avoid inactivation of the reducing agent by metals. Keep in mind that if your sample needs metals for activity or stability that EDTA will keep metals from your sample. 

When working at an alkaline pH, beta-me and TCEP HCl are more stable than DTT. TCEP HCl is more stable at acid pH than DTT. 

DTT reduces nickel ions and can cause problems when purifying His-tagged proteins. To avoid this complication, try beta-me or TCEP HCl as a reducing agent instead of DTT when purifying His-tagged proteins. 

L-cysteine is also a reducing agent. It’s usefulness in crystallization is limited since it likes to form small hexagonal plate shaped crystals. L-cysteine can be a useful crystallization additive, but keep those pesky hexagonal plates in the back of your mind when working with L-cysteine. 

If oxidation of the sample is expected or anticipated, reducing agents should be present during the preparation of the protein (when possible) and should be included, added, or replenished during the final preparation of the sample for crystallization. In vapor diffusion experiments, the reducing agent can be included or added to the crystallization reagent and reservoir.

Preparing the Protein for Crystallization 

Lyophilization 
Avoid lyophilization. Even though there are many examples of proteins which crystallize after lyophilization (lysozyme, thaumatin, hemoglobin), lyophilization is to be avoided when possible. If the protein is lyophilized, it needs to be dialyzed before crystallization. Dialyze the protein against deionized water or a stabilization buffer before crystallization. Dialysis will remove non-volatile buffers and other chemicals which may have been present before lyophilization. 

Ammonium Sulfate Precipitation 
Avoid using ammonium sulfate precipitation as a final purification and/or concentration step. It is often very difficult to completely remove all the ammonium sulfate by a desalting column of dialysis. The remaining trace amounts of ammonium sulfate can interfere with crystallization screening results and create reproducibility problems. It is not uncommon for trace amounts of ammonium sulfate in the sample to cause precipitation or excessive nucleation in screen conditions containing polyethylene glycol and salt. 

Batches 
Avoid combining different purification batches for crystallization trials. Purification conditions and procedures are never identical so each batch should be screened separately. 

Profile the Protein 
Ideally, you will purify your own protein, but this is not always reality. So, it is always a good idea to characterize your protein before beginning crystallization experiments. Profiling your protein before crystallization can often provide valuable clues during screening and optimization of crystallization conditions. Assays to seriously consider: 
• SDS-PAGE 
• Native PAGE or 
• Dynamic Light Scattering 
• IEF (Isoelectric Focusing) Gel 
• Mass Spectroscopy 

The results of these assays can: 
• Determine the purity of the sample 
• Determine the homogeneity of the sample 
• Identify batch-to-batch variations 
• Identify stability problems with the sample 

How Pure? 
How pure should the protein sample be for crystallization trials? As pure as possible. That’s some answer, is it not? Integrating common sense into the question, we might arrive at the following answer. For initial screening, the sample should be at least 90 to 95% pure on a Coomassie stained SDS-PAGE. Finally, it does no harm to screen an “impure sample” as one can always perform further purification. Remember, crystallization used to be considered a very powerful purification tool (and still is!). 
If the initial screen does not produce crystals, any promising results, or it becomes next to impossible to improve crystal quality during optimization, one should consider further purification of the sample. 

Storing the Sample 
Most proteins can be stored successfully at 4°C or -70°C. 

Check with the person preparing the protein or compare your protein to a similar protein in the literature for best storage temperature. Ideally, one should assay the activity and stability of the protein before storage and then later on at various points in time to determine the sample storage stability. Repeated freezing and thawing of the sample should be avoided. Aliquot the sample into multiple small microcentrifuge tubes. Make the aliquots small enough so that the entire aliquot can be consumed in the experiment after thawing. 

Sometimes people like to add glycerol (10 to 50% v/v) to help proteins better tolerate freezing. Avoid this if possible since it is often difficult to remove glycerol by dialysis or filtration. The presence of glycerol is a crystallization variable. Glycerol can behave as a precipitant, an additive, or cryoprotectant and therefore can influence the outcome of a crystallization experiment. 
In general, it is better to store proteins more concentrated than diluted. When too dilute, adsorption of the protein onto the storage container can lead to significant losses. However, precipitation can sometimes be a problem when the protein is stored too concentrated.

Temperature and Crystallization 
Temperature can be a significant variable in the crystallization of biological macromolecules (proteins).(1,5) Temperature often influences nucleation and crystal growth by manipulating the solubility and supersaturation of the sample. Temperature has also been shown to be an important variable with phase separation in detergent solutions during membrane protein crystallization.7 

Control and manipulation of temperature during the screening, optimization and production of crystals is a prerequisite for successful and reproducible crystal growth of proteins with temperature dependent solubility. Christopher et al., testing 30 randomly chosen proteins, found 86% demonstrated a temperature dependent solubility and suggested that temperature induced crystallization could be a generally useful technique.5 Temperature was shown to affect quantity, size, and quality of the crystals as well as sample solubility and preliminary crystallization data. 

One advantage of temperature is that temperature provides precise, quick, and reversible control of relative supersaturation. Using temperature in addition to standard crystallization variables such as sample concentration, reagent composition and concentration, as well as pH can increase the probability of producing crystals as well as uncover new crystallization conditions for a sample. Additional crystallization conditions may uncover reagent formulations more amicable to heavy atom derivatization, cryoprotection, and optimization or at least offer options. Temperature is amenable to control and can be used to carefully manipulate crystal nucleation and growth. This control can also be used to etch or partially dissolve then grow back the crystal in an attempt to improve crystal size, morphology, and quality or assist with seeding. Temperature control is noninvasive and can manipulate sample solubility and crystallization with altering reagent formulation. 

Traditionally, crystallization screens and experiments are performed at room temperature and sometimes 4 degrees Celsius. A reasonable range of temperature to screen and optimize for protein crystallization is 4 to 45 degrees Celsius and some proteins have been crystallized at 60 (glucagon and choriomammotropin) degrees Celsius. A practical strategy would be to screen at 10, 20 (or room temperature) and 30 degrees Celsius when the sample volume permits. Temperature incubations above room temperature should be monitored closely for evaporation from the drop and reservoir. A 2 microliter hanging drop vapor diffusion experiment at 37 degrees Celsius can evaporate in as little as 48 hours depending upon the plate, quality of seal. Microbatch under Paraffin Oil can minimize evaporation problems. In the case of room temperature incubations, temperature control and stability are often minimal since the experiments may be left in the open room. In an open room, temperature fluctuations may be significant, especially over a 24 hour period and on weekends when thermostatic control of the room environment can fluctuate 10 degrees or more. Incubation at 4 degrees Celsius and other temperatures are often more stable since the incubation is performed in some type of incubator. Another source of temperature fluctuation occurs while viewing experiments. The light microscope is a heat source and extended viewing can significantly alter the temperature of small drops. 

Quick efficient viewing can minimize temperature changes. Also, remember to turn off the light source when leaving plates on the stage in one position for more than a few seconds. 

While controlled temperature can be important for consistent results, temperature fluctuation can be useful in obtaining high quality crystals by screening a larger range of crystallization conditions since for a sample with temperature dependent solubility changes in temperature can equate to changes in a crystallization reagent condition.8 Hence, a sparse matrix screen takes on a new dimension when screened at multiple temperatures, or ramped over several different temperatures over a period of time. 

How does one test for the effect of temperature and temperature dependent solubility without consuming a lot of sample? One solution is to set a single crystallization screen at one temperature, allow the experiment to incubate for a week, record the results and then move the plate to another temperature. Allow the experiment to incubate for a week at the new temperature and record the results. If one notices changes in solubility (i.e. clear drop turning to precipitate, or precipitate turning to clear drops) between the two temperatures, then the sample has temperature dependent solubility and temperature should be explored as a crystallization variable. 

Temperature gradients can be used for screening and optimization of proteins with temperature dependent solubility. For screening, set the experiment at one temperature, allow the experiment to equilibrate and then slowly change the temperature to a second temperature. In general, ramp the temperature so that the sample is exposed to an increase in relative supersaturation as the temperature changes over time. In other words, ramp from high to low temperature if the sample is more soluble at high than low temperatures.This can be accomplished using a programmable temperature incubator. A temperature gradient or ramp, allows one to slowly approach temperatures where a sample may have a decrease in solubility with a corresponding increase in relative supersaturation. Published examples of temperature gradient or temperature ramp crystallization include elastase (25 to 20 degrees Celsius gradient), alpha-amylase (25 to 12 degrees Celsius gradient), and insulin (50 to 25 degrees Celsius gradient).(9,10,11) 

To demonstrate how screening temperature could affect and enhance the results obtained from a preliminary crystallization screen, a programmable temperature incubator was used to screen 4 different temperatures. Using Glucose Isomerase and Crystal Screen, sitting drop vapor diffusion experiments were set using Cryschem plates at 4, 15, 25, and 37 degrees Celsius. Drops were observed daily and the results were quite interesting. Glucose Isomerase crystallized in 19 conditions at 25 degrees Celsius, 23 conditions at 15 degrees Celsius, 28 conditions at 4 degrees Celsius, and 12 conditions at 37 degrees Celsius. A similar approach with Trypsin, yielded crystals in 8 conditions at 15 degrees Celsius, 4 conditions at 25 degrees Celsius, and 7 conditions at 32 degrees Celsius. In the case of Trypsin, a single set of Cryschem plates were set and the plates simply moved from one temperature to another over a period of a weeks time, scoring results before each temperature change. 
Solutions for Crystal Growth 

Temperature Tips 
• For proteins with “normal” solubility, in high salt the protein will be more soluble at cold than at warm temperatures. 
• For proteins with “normal” solubility, in low salt the protein will be more soluble at warm than at cold temperatures. 
• Proteins with “normal” solubility will precipitate or crystallize from lower concentration of PEG, MPD, or organic solvent more slowly at low than at high temperatures. 
• Diffusion rates are less and equilibration occurs more slowly at low than at high temperatures. Crystallization may occur more slowly at low than at high temperatures. 
• Temperature effects can be more pronounced at low ionic strength reagent 
conditions. 
• Do not use the appearance or non-appearance of crystals at various temperatures to gauge the effectiveness of temperature as a crystallization variable. Rather, use the difference in the solubility at different temperatures to gauge the effect temperature has on sample solubility. If an effect is observed, explore temperature as a crystallization variable. 
• Temperature can effect different crystal forms and growth mechanisms.12 
• When incubating experiments below and above room temperature and viewing experiments at room temperature, condensation can be a problem. To minimize and avoid condensation with vapor diffusion experiments, stack a “Dummy Plate” with reservoir filled with water and sealed, at the bottom and top of the stack of plates. This will slow the temperature change in the sandwiched plates and minimize condensation. 
• The Microbatch method works well for temperature exploration. In a traditional Microbatch experiment, the relative supersaturation of the system does not change since, in theory there is no vapor diffusion. However, if the sample exhibits temperature dependent solubility, temperature can be used to manipulate sample solubility in a Microbatch experiment. Another plus of using Microbatch is the lack of condensation while viewing the experiment. 
• Condensation with a hanging drop can mean alteration of your drop with the when the condensation mixes with the drop. Condensation with a sitting drop can mean there will be no mixing of the condensation with your drop, unless the condensation falls into the drop. Moral, sitting drop has less change for mixing with condensation. 
• To dry up condensation, add a small amount of concentrated salt solution to the reservoir. Keep in mind this might also dry your drop a bit. 
• Nucleic acid temperature stability allows one to examine temperatures between 4 and 35 degrees Celsius. 
• Ideally, one should set the experiment at the eventual incubation temperature and all reagents, samples, and plates should be equilibrated to the incubation temperature. This is a reality for room temperature setups and 4 degrees Celsius setups for those of us with cold rooms. For the rest of us, we can set the experiment at room temp and then toss it into the incubator. Or, for 4 degrees Celsius set ups, one can cheat. Simply incubate the reagents, sample, plates and slides in the refrigerator before set up. During the set up, place materials in a tray full of ice. Maintain the plates on ice during the set up. Seal and move smartly to the 4 degrees Celsius incubator. 
• Increasing temperature increases the disorder of reagent molecules. Varying the temperature of a crystallization experiment can manipulate sample-sample as well as sample-reagent and reagent-reagent interactions. Such manipulations may have an impact on interactions which control nucleation and crystal growth. In addition, such interactions may have an impact on crystal packing as well as the termination of crystal growth. Hence, temperature can impact nucleation, growth, packing, and termination. 
• Temperature can be a habit modifier and change the crystal lattice. For example, at temperatures below 25 degrees Celsius and in the presence of sodium chloride and acidic pH, the tetragonal form of lysozyme is favored. Under similar reagent conditions above 25 degrees Celsius, the orthorhombic form is favored.13 
• The preparation of heavy atom isomorphous derivatives can depend upon the temperature of the experiment. In most cases, it seems the soak temperature is the same as the crystallization temperature. 

References 
1. Giege, R., and Mikol, V., Trends in Biotechnology (1989) 7, 277. 
2. McPherson, A., European J. Biochemistry (1990) 189, 1. 
3. A. Ducruix and R. Giege, Editors, Crystallization of Nucleic Acids and Proteins: A Practical Approach, IRL Press at Oxford University Press, 1991. 
4. Lorber, B., and Giege, R., Journal of Crystal Growth (1992) 122, 168-175. 
5. Christopher, G.K., Phipps, A.G., and Gray, R.J., Journal of Crystal Growth (1998) 191, 820-826. 
6. Haser, R., et al., Journal of Crystal Growth (1992) 123, 109-120. 
7. Garavito, R.M., and Picot, D., Journal of Crystal Growth (1991) 110, 89. 
8. Drenth, J., Crystal Growth (1988) 90, 368. 
9. Shotton, D.M., Hartley, B.S., Camerman, H., Hofmann, T., Nyborg, S.C., and Rao, L., Journal of Molecular Biology (1968) 32, 155-156. 
10. McPherson, A., and Rich, A., Biochem. Biophys. Acta (1972) 285, 493-497. 
11. T.L. Blundell, and L.N. Johnson, Protein Crystallography, Academic Press (New York) 1976, 59-82 (method by Guy Dodson). 
12. A. McPherson, Crystallization of Biological Macromolecules, Cold Spring Harbor Laboratory Press, 1999. 
13. Ataka, M., and Tanaka, S., Biopolymers (1986) 25, 337.

Molarity 
Molarity is the number of moles of solute per liter and in represented as M, such as 3.0 M ammonium sulfate. Molarity is the ratio between the moles of dissolved solute (solid stuff) and the volume of solution (liquid stuff) in liters. The accepted volume of the solution is 1 L, so a 1M (molar) solution would be 1M = 1 mole of solute/1 L solution. Molarity is a way of determining the concentration of a solution. Dilute solutions are typically expressed in terms of millimolarity (mM) where 1 mM = 0.001 M. Typically, in crystallization we are asked to make something like a 3.5 M solution of ammonium sulfate. To do this we need to know the molecular weight (Mr) of ammonium sulfate (132.14 g/mole), the volume of solution to make (let’s make 500 ml or 0.5 L), and the desired concentration (3.5 M). Then we calculate: 

# grams required = (desired Molarity)(formulation volume in liters)(Mr) 
# grams required = (3.5 mole/liter)(0.5 liter)(132.14 g/mole) = 231.25 g 

To formulate the 3.5 M ammonium sulfate we then weigh 231.25 g of ammonium sulfate and add deionized water to dissolve the ammonium sulfate and then adjust the final volume to 0.5 liter (500 ml). Do not simply add 500 ml of water to 231.25 g of ammonium sulfate. Ideally one should use the most precise measuring instrument possible such as a volumetric flask. A less desirable instrument would be a graduated cylinder and the least desirable would be a beaker. Molarity is typically used as a concentration unit for salts 1,6-hexanediol, detergents, and some additives. 

% w/v 
% w/v (percent weight/volume) is often used when formulating high molecular weight polyethylene glycols which are typically solids as well as some additives in solid form. % w/v is the weight of a solute in a given volume. % w/v = gram per 100 ml. For example, let’s make 500 ml of a 50% w/v PEG 4000. 

# grams required = (desired concentration in g/100ml)(formulation volume in milliliters) 
#grams required = (50 g/100ml)(1,000 ml) = 500 grams 

50% w/v PEG 3350 is 500 g of PEG 3350 in a final volume of 1,000 ml and not 500 g of PEG 3350 plus 500 g of water. To make 1 liter of a 50% w/v solution of PEG 3350, weigh 500 grams of PEG 3350 into a volumetric flask and bring the final volume to 1 liter with water. Do not make the mistake of adding 500 grams of PEG 3350 to 500 ml of water and believing you have made a 50% w/v solution. 

% v/v 
%v/v (percent volume/volume) is often used when formulating liquid, low molecular weight polymers (PEG 400), organics (MPD), and organic solvents (iso-propanol) into stock solutions. % v/v is the volume of a solute in a given volume and is not a volume plus a volume. For example, a 50% v/v iso-propanol is 50 ml of iso-propanol brought to a final volume of 100 ml with water and is not 50 ml of iso-propanol plus 50 ml of water. Formulating reagents using %v/v can be done more precisely and accurately using the mass of the solute being formulated instead of the volume of the solute. To do this we need to know the density of the solute. For example, let’s formulate 500 ml of 30% v/v (±)-2-Methyl-2,4-pentanediol (MPD). The density of MPD is 0.925 g/ml at 25° Celsius. 

#grams required = (desired concentration in ml/100ml)(density of solute)(formulation volume in milliliters) 
#grams required = (30 ml/100ml)(0.925 g/ml)(500 ml) = 138.75 grams 

30% of 500 ml is (0.3)(500) = 150 ml. To account for the difference in density of MPD we multiply 150 ml by 0.925 g/ml and obtain 138.75 grams. To formulate a 30% v/v MPD solution by mass we would then add 138.75 grams of MPD to a volumetric flask and bring the final volume to 500 ml at 25° Celsius with deionized water. 

% saturation 
% saturation is the concentration of material in solution as a percent of the maximum concentration possible at the given temperature. A saturated solution is one where there is equilibrium between undissolved solute and dissolved solute. To make a saturated solution, a salt is added to water and often warmed to enhance solubilization. Complete dissolution is desired. Upon cooling, some of the solute (salt) will crystallize out and leave behind a saturated solution. The actual concentration of a saturated stock depends upon the temperature of the solution. For example, at 0°C, 127.5 g of potassium iodide can be dissolved into 100 ml of water, but at 20°C, 144 g of potassium iodide can be dissolved into 100 ml of water. Therefore, depending upon whether the solution is kept at room temperature or in the cold, the concentration can be very different. % saturation is a rather old school way to make salt solutions for crystallization. However, since we often perform crystallization at different temperatures, the actual concentration in the bottle, reservoir, or drop can be very different. Plus, exact reproduction of a stock solution not only depends upon careful mass and volume measurement, but also temperature. Keep life simple, avoid reproducibility problems and stick with M, % w/v and % v/v when formulating your solutions. 

Mg/ml 
Mg/ml (mg/ml) is typically used to express or determine protein concentration. To make a 20 mg/ml lysozyme solution we would weigh 20 mg of lysozyme and simply add 1 ml of buffer (20 mg plus 1 ml). However, others might weigh 20 mg of lysozyme and add 980 µl (0.98 ml) (20 mg in 1 ml). Be sure you document which method was used to avoid a slight concentration inconsistency and potential reproducibility problems later.

The role of water in protein crystallization is significant as protein crystals are highly hydrated. The amount of water in a drop or a crystal, the purity, the chemical composition, and how the water interacts with itself, the sample, and reagents are all critical variables in a protein crystallization experiment. Consistency is typically a good thing to keep in mind when trying to reproduce crystallization results. Being consistent with one’s treatment, handling and use of water can be an important crystallization variable. Consider the source of water. One should use pure and fresh water for crystallization experiments. Type 1+ ultrapure grade water with a resistivity of greater than 18.2 megohm, conductivity of 0.055 microsiemens or less, a total organic carbon (TOC) of less than 5 ppb, bacterial count of less than 1 CFU/ml, and particulates (size > 0.22 µm of less than 1/ml is a good source of water for protein crystallization. Purchasing and using a quality water purification system is one part of the water source equation. Different water systems and different methods of water purification produce different specs of water. For example, glass distilled water typically has a pH of 5.5 while reverse osmosis and deionization systems typically produce water with a pH of 7. 

Another part of the water equation is the proper use and maintenance of the water purification system. Carefully follow the manufacturer’s use, maintenance, and service guidelines. Establish regular quality control practices to ensure the consistent quality of the water. Using a water purification cartridge beyond its expiration date or failing to clean the system on a regular basis can lead to water with higher resistivity, higher conductivity, as well as microbial contamination. Microbial contamination can change the pH of the water as well as expose your sample to proteolysis. Use fresh water. Purified water is deionized and can leach zinc, lead, copper, iron, aluminum and other substances from glass or plastic storage containers. Purified water stored in plastic containers for several weeks has nearly the same level of TOC as tap water. 

Use fresh, pure water for your crystallization experiments. When having trouble reproducing experiments, consider water, the difference in purity and the difference in source as potential crystallization variables. 

Hampton Research uses Type 1+ ultrapure water: 18.2 megaohm-cm resistivity at 25°C, < 5 ppb Total Organic Carbon, bacteria free (<1 Bacteria (CFU/ml)), pyrogen free (<0.03 Endotoxin (EU/ml)), RNase-free (< 0.01 ng/mL) and DNase-free (< 4 pg/µL), including but not limited to crystallization screens, Custom Shop reagents, and Optimize buffers, salts, and polymers.

Xenon is a noble gas that binds to specific sites in a macromolecule. Xenon-protein complexes can often serve as heavy atom complexes for MIR structure determination. Heavy atom derivatives of protein crystals can be produced by pressurizing native crystals with xenon gas. In the vast majority of cases, the modification of the mother liquor to determine soaking conditions is avoided since the crystal and mother liquor are simply placed in a chamber pressurized with xenon gas. Another advantage of working with xenon is that it interacts weakly with a protein so isomorphism of the derivative with the native crystal is high. Also, the number of binding sites as well as the binding occupancies can often be changed by altering the pressure of the xenon gas. Xenon binding sites often differ from heavy metal binding sites so it can be useful to try xenon when traditional heavy atom soaks fail. Xenon binding is often reversible so if one has very few crystals, the same crystal could be used for heavy atom soak. 

The interaction between xenon and the protein is typically confined to weak dispersion forces and does not involve electrostatic interactions. Therefore, xenon should bind to different locations of the protein compared to most other heavy atom compounds. The weakness of the interaction between xenon and the protein makes it unlikely that xenon will interfere with crystal contacts. Xenon derivatives usually show good isomorphicity with the native crystal, resulting in high phasing power. 

Xenon seems to have little or no effect on the pH or ionic strength of the mother liquor. The weakness of the interactions between xenon and proteins requires a significant xenon concentration in the crystal’s mother liquor to enforce sufficient occupation of a potential binding site so most uses of xenon reported in the literature make use of high pressure equipment for xenon derivatization. At this time is appears cryocooling does not alter the protein’s xenon binding properties. 
Xenon derivatization can be performed at room temperature and data collection with xenon derivatives can be performed at room temperature using specially made pressure cells and capillaries or at cryogenic temperatures using CryoLoops. 

Another application of xenon in protein crystals is the crystallographic imaging of disordered areas of lipids or detergents in crystals of membrane proteins. 

References for using xenon as a derivative: 
1. Binding of Xenon to Sperm Whale Myoglobin. Schoenborn B.P.; Watson, H.C.; Kendrew, J.C. (1965). Nature, 207, 28-30. 
2. Cavities in proteins: structure of a metmyoglobin-xenon complex solved to 1.9&Angring. Tilton, R.F.; Kuntz, L.D.; Pesko, G.A. (1984) Biochemistry 23. 2849-2857. 
3. Using Xenon as a Heavy Atom for Determining Phases in Sperm Whale Metmyoglobin. Vitali, J.; Robbins, A.H.; Almo, S.C.; Tilton, R.F. (1991). Journal of Applied Crystallography, 24, 931-935. 
4. On the Preparation and X-ray Data Collection of Isomorphous Xenon Derivatives. Schiltz, M.; Prange, T.; Fourme, R. (1994). Journal of Applied Crystallography, 27, 950-960. 
5. Successful flash-cooling of xenon-derivatised myoglobin crystals. Soltis, S.M.; Stowell, M.H.B.; Wiener, M.C.; Philips, G.N.; Rees, D.C. (1997).Journal of Applied Crystallography, 30, 190-194. 
6. Freeze-Trapping Isomorphous Xenon Derivatives of Protein Crystals. Sauer, O.; Schmidt, A.; Kratky, C. (1997). Journal of Applied Crystallography, 30, 476-486. 
7. Protein Crystallography at Ultra-Short Wavelengths: Feasibility Study of Anomalous Dispersion Experiments at the Xenon K-Edge. Schiltz, M., Kvick, A., Svensson, O., Shepard, W., De LaFortelle, E., Prange, T., Kahn, R. & Fourme, R. (1997). Journal of Synchrotron Radiation , 4, 287-297. 
8. A method to stabilize reduced and/or gas treated protein crystals by flash cooling under a controlled atmospher 
9. Xavier Vermede et.al. J. App. Cryst, (1999) 32(3) 505-509.

Protein samples are yummy treats for microbes such as bacteria, yeast and fungus which secrete proteases that like to chop your sample into tasty bites, ruining your crystallization setup. These same microbial menaces also like to dine on polymer based crystallization reagents such a polyethylene glycols and even buffers. So even if the bugs and their secreted proteases do not chop your sample into bits, they can degrade reagents, alter the pH of the solution, or generate chemical species which can influence crystallization and even make reproducing conditions a pain in the tush. To prevent or at least minimize your sample and reagents from becoming Sunday Crystallization Brunch for microbes, consider the following suggestions: 

Sterile filter the sample into a sterile tube using a 0.2 micron filter before setup to remove microbes. Do not leave the sample on the bench at room temperature for extended periods. Store unused sample appropriately (4, -20, or -80°C; only you know best). Those of you working with engineers, physicists, computer scientists, or folks from outside the life science field, take a moment to realize that quarks, gluons, hard drives and other physical things these folks are used to handling do not support microbial growth. Take a little time to help these crystallization rookies understand what happens to samples left on the bench for a day or three and the significance of keeping things clean to prevent microbial (as well as chemical) contamination. Case in point – Caller: “My lysozyme stock that was growing crystals no longer grows crystals.” Tech Support: “Where are you storing the lysozyme when it is not being used?” Caller: “Same place as the reagents.” Tech: “Which is?” Caller: “On the bench (i.e. room temperature).” Tech Support: “For how long?” Caller: “Oh, not long, maybe a few days, a couple weeks at the most”. Hey, nothing wrong with these types of questions. We all gotta learn some time. Just be sure and help that engineer, computer scientist and new post doc from a physics lab who is helping you with your new AFM machine or testing the high throughput robot, to understand proteins are food for microbes. Who knows, maybe they will help you with the next Windows update installation. 

Sterile filter water into a sterile container before formulating crystallization reagents that cannot or should not be filtered (detergents, gels, and some high molecular weight polymeric agents too big for filtering). 

Sterile filter (0.2 micron pore size) buffers, salts, polymers, and diluted organics into sterile containers. Sterile containers such as polypropylene or PETG plastic are readily available today and are cost-effective, time-saving alternatives to glass and autoclaving. 
When using a stock that has been sitting about for more than a couple of weeks, give the bottle a swirl and look for signs of microbial growth such as a settle, faint white, off white or yellow to brown precipitate. Sometimes slightly precipitated salts will resemble microbial growth to the untrained eye. To help differentiate precipitate from microbial growth, try warming the solution in your hands for 5 to 10 minutes. If the material disappears it might well be precipitate. Microbial growth will not dissolve. Although we are not endorsing the sniffing of reagents, especially since some crystallization reagents can be hazardous, precipitates will not smell, while microbial growth will often stink. 

Use sterile pipet tips when pipetting reagents, sample, and water into plates. Keep your filthy paws away from your pipet tips and do not touch pipet tips to the counter, your lab partner or other non-sterile items. If you set your pipet down it had better not have a tip on it or the tip will likely touch something non-sterile. 

Keep crystallization plates in their sealed wrappers until just before use. This will prevent airborne microbes from setting up home in your crystallization plates.

Zero or “0” is often used to represent a clear drop when scoring crystallization experiments. While on the subject of zero or nothing, consider using -1 or some other more interesting and exciting score to represent a non-experiment. A non-experiment is a drop that has fallen off or a drop that was never pipetted (human or robot error). Better to use a non-experiment score such as -1 than a “0” since someone reading your scores might mistake a “0” as a clear drop.

Microvolume crystallization by dialysis was first reported by Zeppenzauer in 1971 (Methods in Enzymology, ibid ref. 1. Vol. 22, page 253). The Zeppenzauer cells are homemade from capillaries with the ends of the tubes either covered with dialysis membrane or plugged with acrylamide, silica, or agarose gel. If covering the tube with a dialysis membrane, one can use small o-rings to secure the membrane over the tube. The capillary is then placed in an appropriate chamber with a volume of reagent sufficient to cover the ends of the tube.

1 to 5 mM zinc (most frequently as zinc chloride or zinc sulfate) can be a useful crystallization additive and typically reduces the solubility of the sample. Zinc is not very soluble and can readily precipitate or crystallize out of reagents as higher concentrations.

If you know the Matthews volume, then you can convert to protein concentration with: 

[protein] (mg/mL) = 1/( V_M (A^3/Da) ) * 1.66e-21 (mg/Da) * 1e24 (A^3/mL) 

For example, lysozyme crystals (V_M = 2.0 A^3/Da) contain about 830 mg/ml of protein. Most protein crystals are in this ballpark (because V_M doesn't have much of a range). 

If you want molarity, then you also need to know the molecular weight (MW): 

[protein] (mol/L) = [protein] (mg/mL) / MW (g/mol) 

Or about 60 mM lysozyme monomers in a crystal. 

Reference: Ccp4bb, December 13, 2010. James Holton, Lawrence Berkeley National Laboratory, USA. 

Or 

c = Z/(NxV) 

where Z=number of molecules per unit cell, N=Avogadro's number, V=volume of the unit cell in liter, c=concentration in molar 

Reference: Ccp4bb, December 13, 2010. Filip Van Petegem. 

Specific density of protein is about 1.47 g/cm3. 

Accurate calculation of the density of proteins. M. L. Quillin and B. W. Matthews. Acta Cryst. (2000). D56, 791-794 [doi:10.1107/S090744490000679X ]

Unidentified blobs and bits in a crystallographic structure are often times something in the crystallization reagent or sample buffer or cryoprotectant. To help identify what the unidentified blob or bit might be, think about everything the protein has come into contact with since its inception. It would something from inside the cell, something during the expression, refolding, purification or crystallization and cryoprotection. It could be a buffer, a ligand, or a detergent carried through several steps of purification. If a polyethylene glycol (PEG) was used in the crystallization, it could be PEG. Proteins can be gregarious sponge like molecules and can cling to spurious small, medium and large molecules. 

Ask yourself the following. What is in the crystallization solution? It would be rare that you would observe something that was not added during crystallization or purification. What are the chemical interactions with the unknown density? This can give some clue as to the nature of the ligand. Anions, for example, will normally be interacting with H-bond donors and positively charged side chains. What does the anomalous map look like? This can be useful if you suspect the presence of heavy atom scatterers, such as sulfur atoms in sulfate or iodine atoms as iodide.

For a crystal to grow, the system must be supersaturated, in non-equilibrium. But first a stable nucleus must form. A stable nucleus is an aggregate of the macromolecule. A stable nucleus is an aggregate of such size, shape, and properties that it will enlist new molecules into it's growing surface faster than others are lost into solution.

Source: Worthington catalog number LS001432 for 100 milligrams. Code: CDTLCK 3X crystallized and treated with 1-chloro-3-tosylamido-7-amino-2-heptanone (TLCK) to inhibit trypsin activity [Shaw, et al., Biochemistry, 4, 2219 (1965)]. Dialyzed against 1 mM HCl to remove autolysis products and low molecular weight contaminants. Supplied as a dialyzed, lyophilized powder. Store at 2-8°C. Source: Bovine Pancreas 
Minimum Activity: =45 units per mg protein 

Crystallizes in Hampton Research Index crystllization screen reagents 66/F6, 68/F8, 69/F9, 86/H2 and 88/H4. Recommended protein concentration 20 mg/ml in 0.05 M Hepes pH 7.0.

Hemoglobin (horse) 
Source: Sigma H4632 

Crystallizes in Index D1, D3, D5, D6, D7, D8, D9, D10, D11, F1, F2, F3, F8, F9, F10, F11, F12, G1, G2, G3, G4, G5, G6, G7, G8, G9, G10, G11, H2, H3, H4, H5, H6. H7, H8, H11, H12. 

Protein concentration 20 mg/ml in 50 mM Hepes pH 7.0.

Filtering the protein 
To reduce the number of crystals and increase their size, try filtering the protein solution prior to setting up the experiment. Try the following filter sizes: 0.22, 0.1 micron and 300 kD molecular weight cut-off. Try the Millipore centrifugal filters. 
Naomi Chayen 

Pseudo-Microseeding 
When working with crystals that grow fairly rapidly (one day) try the following. Pipette multiple protein drops (2 to 4 works best) onto the cover slide. Using a single pipette tip, get the reservoir solution to mix with the first drop. Now, go back to into the reservoir with the same tip to get the reservoir solution for the second drop. Continue for the remaining drops with the same tip. In certain cases, seeding starts very quickly, so by using the same tip one can introduce minute seeds to successive drops. Use the same cover slide with multiple drops to minimize evaporation. 
Mike Sintchak 

Low molecular weight PEGs 
When screening with low molecular weight PEGs try microbatch. Crystals appear rapidly with PEG 400-2000. To convert to vapor diffusion use 0.2 M buffer in the well and a 1:1 drop ratio. Try using a positive displacement pipette such as the Anachem Microman 1 - 10 microliter. These are much more accurate. 
Lesley Haire 

Purest is not the best 
A protein which was purified and showed some faint bands of contaminants on a native gel was crystallized and solved successfully. The same protein, purified by HPLC and resulting in a single band native gel did not crystallize. 
Michal Harel 

Cryoprotectant 
When making up cryoprotectant solutions containing glycerol, put a test tube of glycerol in a beaker of warm water. The viscosity falls and it is easier to pipette accurately. 
Elspeth Garman 

Mass spectrometry 
We found mass spectrometry like ESMS and MALDI highly efficient in determining impurities and/or microheterogeneities in our protein sample/batch. In most cases it is a simple, straight forward method which requires a minimum amount of sample. In some cases it has shown to detect impurities/microheterogeneities when other techniques did not. 
David Leys

Shape of Drop 
One of my proteins produced only zillions of tiny useless crystals. When I mixed the drops the conventional way-mixing well, overlaying, etc. the protein with precipitant solution. Large, gorgeous crystals were produced when I crossed the drop, creating a gradient within the drop. This worked best, setting up sitting drops with vapor diffusion.
Ursula Kamlott 

Don't Flip 
When removing crystals from a hanging drop, I sometimes find that the biggest crystals fall against the coverslip and are impossible to resuspend without damage. I had our glass shop make a stand to transfer the coverslip to that enabled me to manipulate the crystals more easily. 
Dennise Dombroski 

Extreme Soak 
For soaking crystals with compounds with limited solubility I have tried two "extreme ways" (although not much- and more experiments should be tried): 
- Leave some solid compound in the soaking solution. 
- Dissolve some compound in n-octanol and layer the octanol solution on top of the soaking experiment. 
These two provide (hopefully) more or less constant concentration of the compound in the soaking solution. The octanol layer may help reduce air oxidation by preventing direct contact of soaking solution to air. 
I found that 200 µl of soaking solution in a well of the 24 well Linbro plate is a good volume to work with: 
--Not too little that possibly causes concentration change due to evaporation (Sealed well) and not too much that the crystals get lost in the solution. 
Jirundon Yuvaniyama 

The Glycerol Effect 
Glycerol has many benefits but also some drawbacks. We found it to be beneficial with one protein we were working with; this protein is a transpeptidase called Mur A. The protein is quite soluble and could concentrate to 20 mg/ml but it lost activity over time when stored at ­80°C. We therefore dialysed it into 50% v/v Glycerol to see if the activity could be retained for longer, this would allow us to make a large batch rather than regular smaller batches, for crystallisation. On dialysis we found a significant reduction in the volume of protein, so much so that the protein had concentrated from 5-10 mg/ml to almost 50 mg/ml. Activity was also found to be retained with no significant loss after 6 months at ­80°C. This gave us a method for storing large batches of Mur A at ­80°C, without losing activity and also resulted in a sample pre-concentrated for crystallisation and containing a cryo-protectant. Other sugars gave similar effects; sucrose, sorbitol, etc., but none were as effective as glycerol in achieving the 50 mg/ml final concentration. 
Anil Mistry 

Iodoacetic Acid 
Add a small amount of (~ 1%) iodoacetic acid to buffer solutions. This helps prevent aggregation by carboxymethylation of cys. Also, iodoacetic acid seems to help form salt bridges and aid in crystallization. 
Bernie Santarsiero 

Soaking Crystals to Improve Resolution 
Try soaking poorly diffracting crystals in higher concentration of precipitate‹ammonium sulfate or PEG. It may take several weeks so test after 1 or 2 months. 
Irene Weber 

Rapid Preliminary Screening of Protein for Aggregation Using Protein Quantities 
When only small amounts of protein are available, it is not feasible to screen many compounds which promote monodispersim using a dynamic light scattering machine. To detect aggregation, we use a pseudo-native gel approach: 1l of protein is mixed with 1 l additive from Hampton Additive/ or Detergent Screens. These samples are then incubated at room temperature for 20-30 minutes and then placed in 2x sample buffer containing no DTT, no SDS and are not boiled. These samples are run on a standard SDS-PAGE gel. We have screened many additives using this approach and it has given us leads for subsequent optimization of protein buffers. 
Tom Zarembinski 

HPLC Profile 
Keep an HPLC (Reverse phase) profile of your protein before crystallisation and after crystal formation. It can be used as a quality control and tells you if any modifications have occurred. 
Glenn Dale 

Don't Throw Away Without Looking Close 
Look closely at your old test tubes when cleaning the place. Proteins do crystallize on the walls of the tube when stored in a cold room. I picked up my tubes from the wastebasket and an X-ray was made from an old supersaturated protein tube. 
Kalevi Visuri 

Concentration of Protein without Aggregation 
Use centipreps (millipore) for concentrating protein >O.5 ml. Protein concentrating away from membrane. (No micro high concentration, ppt on membrane), works very nicely for a number of proteins. 
Paul Reichert 

Preserve Hampton Solutions 
To preserve Hampton solution when you or a co-worker gets a "hit" and suspect the Hampton solution may be "magic" or you cannot reproduce crystals with lab-made solution‹make the reservoir solution from lab ingredients and use your homemade solution for reservoirs. Use the Hampton "magic" solution only for the drops thus using a few ul per experiment rather than O.5 ml. Saves having a 48 well screening solution set with one tube empty and the rest at 8 ml and still allows the superstitions or in Alex's case, the contaminated solution to reproduce crystals. 
Cheryl Janson 

Recycle Your Precipitate 
If your protein refolding reaction has a low yield and produces lots of precipitate try collecting the precipitate, resolubilize in GuHCl, and refold again. This material is sometimes more pure than the washed inclusion bodies. 

Neali Armstrong 
Columbia University: naa15@columbia.edu 
90% Solutions 
When optimizing or making solutions for a random scan, omit the buffer (@ final O.1 M) so that your stock is 90%. Prior to putting your stock in the well, add 100 uL of 1M buffer of your choice. Next, add 900 uL of 90% stock and mix. This reduces the number of tubes for crystallization (precipitant) stocks and allows flexibility in buffer identity and pH range. Be sure to make plenty of (~20mL) of precipitant, you¹ll need 900 uL per buffer. 
Anna Stevens 

Check Both Liquid Nitrogen + Stream Flash Cooling 
If your crystal does not freeze well in the cold stream try liquid nitrogen or vice versa. 
Hans Parge 

Mounting Needles 
Make a thin needle out of a glass capillary. Fish out crystal needles from drop with the capillary by holding the capillary parallel to the crystal. This will pick up less mother liquor (reduce background) and put less stress on the crystal. 
Frederick de Mare 

To Determine the Optimal Concentration of Your Protein for Screen I and Screen II 
Try (set up) drop no. 6 first (of Screen I). It should produce light precipitation‹if ppt appears too heavy, reduce the protein concentration by 1/2 and try again. If there is no precipitation in drop 6 try drop 4. If there is no precipitation in either of the drops concentrate the protein 2 fold and try again. 
Jaru Jancarik 

Cryoprotectant Additive 
I tried adding 1-10 mg/ml BSA in the cryoprotectant when soaking crystals that would crack. The one time it was used, the crystal did not crack and froze nicely. I don¹t know if the BSA was the reason for successful freezing. I was wondering if anyone else has tried this. 
Laura Pelletier 

Slow Cryosoaks for Improved Crystal Stability 
In a sitting drop well, cryoprotectant (20% Glycerol in crystallization buffer) should be dribbled down the side of the well in the following manner: See drawing. 
Bryan Prince & Melissa Harris 

Crosslinking of Crystals 
When crystals are fragile or you want to transfer them to a different mother liquor e.g. for heavy atom soaking, why not try crosslinking your crystals with 0.1% glutaraldehyde in your mother liquor. This can be done by adding the glutaraldehyde directly to the drop or placing it next to the drop and allowing for vapour diffusion. This crosslinking enabled us to solve Mod A at 1.2 A resolution. 
Clare Stevenson 

Know when enough is enough! 
As a graduate student, I spent countless hours and quantities of protein trying to get crystals of a single construct. We never got crystals. Another group got the structure of a homologous protein that was auto-digestive. Has we stopped after 400 trials and altered our construct, perhaps we would have faired betterŠ we couldn¹t have faired any worse. There is a reasonable statistical argument to demonstrate that 400 trials are a good limit. 
Brent Segelke 

Buffer Screening 
To find the best buffer system which will keep your protein happy, stable, and soluble for concentration prior to crystallisation, setup your protein (at 1-2 mg/ml) in hanging drops over 1 ml well solutions containing a series of buffers at various pH¹s, with various additives/stabilizers, etc. (but no precipitant!). Checking for drops which are clear will give an idea of which solutions keep the protein happy. In addition, as the system attains equilibrium some in situ concentration of protein will be induced due to the bulk difference between the drop and well volumes, hence an idea of how the protein behaves upon concentration in this solution will also be observed. Modifying a clear or slightly clear drop by adding a higher concentration of buffer to the well may even produce crystals. However, the main piece of information this method can produce is an idea of which buffer system and additives to put your protein into prior to concentration a crystallisation screening. 
Anil Mistry 

Soft Crystals 
If you have crystals that are very sensitive to being touched (they break) or stick to the glass or dish, use a pointed strip of parafilm to move them. Otherwise, grow the crystals on parafilm and punch "wells" around the crystals to move it. 
Allan D'Arcy 

Crystal Annealing 
As I said in my talk, give it a go. You might be surprised!! 
Clare Stevenson 

No More 4 Degrees Celsius 
Prepare trays for crystallisation, leave at 4°C, and fill polystyrene box or flat container with ice. Imbed a metal plate in the ice and set out cover slips. When you¹re ready to set out crystallisations place the trays in the bed of the ice and prepare drops, when finished transfer to 4°C. Simple but it works. 
Marie Anderson 

Floating Boat in the Sea - Right Side Up or Down? 
Single dehydrant/reservoir Hanging Drop/ Sitting Drop vapor diffusion using microbatch plate in suitable container. 
Application for Hampton Screening: 
- Low Ionic Strength Screen 
- All Additive Screens 
- Nucleic Acid Mini Screen 
- All Detergent Screens 
Betty Yu 

Cross Seeding to Generate Crystals of a Related Protein or Protein/Inhibitor Complex 
If your protein or protein complex fails to crystallize try seeding from crystals of the same protein (if it's a protein you want to crystallize) or a related protein for apo protein crystals. Serial micro seeding works best, make sure you look at the drops after seeding to identify any visible crystal seeds in there (i.e. not new crystals) 
Margaret O'Gara 

Advice for the Follically - Challenged Crystallographer 
It has been long accepted that crystallizers with an abundance of facial hair have highly successful careers (Leeds University‹personal observation). This was thought to stem from the fact that matter found its way from the hair into the trials and acted as a nucleation centre-BUT SERIOUSLY; Addition of small grains of sand to a crystallization drop that is "close to producing" crystals can aid in nucleation and crystal growth. 
Jonathon Hadden 

Raise the DMSO? 
If higher DMSO does not damage your protein, try higher concentrations of DMSO (10-20%) for crystallization. It can help when dealing with insoluble compounds and is an excellent cryo-protectant. Freeze directly from drop! 
Melissa Harris 

TCEP as a Reducing Agent 
Use TCEP instead of DTT as a reducing agent in your protein solution. It isn¹t oxidized as quickly as DTT. Be sure to watch the pH of your solutions because it is very acidic. 
Barbara Brandhuber 

Temperature Variation 
To grow crystals at different temperatures around room temperature search the lab for spots that are consistently at higher or lower temperatures. A difference of several degrees can be found. Temperature shifts can be easily made by moving crystals to a different place. (Check office shelves too!) [Discovered by Charles Reed in my lab] 
Irene Weber 

Drop Drying Technique (courtesy of Ron Rubin in our lab at Parke Davis) 
When setting up drops with a protein which has low solubility a low starting concentration has to be used. Setup larger drops (5-10 ul) and leave them to stand "dry" for 3-5 minutes at room temperature /4°C prior to inverting over a well of a Linbro plate, this should allow some pre-concentration. 
Anil Mistry 

Using DLS to Test for Irreversible Aggregation 
Generally, people concentrate a protein to 10 mg/ml or higher, then dilute 2-fold when setting up their hanging drops by doing a 1:1 minx. What if in concentrating to 10 mg/ml aggregation has been induced which is irreversible, such when pipetting your 1:1 mix hanging drop, it already contains aggregates‹a bad starting point. 
(For monodisperse protein samples) 
Using DLS test the limit of concentrated protein, i.e., the maximum concentration that can be achieved before a polydisperse signal is obtained. Then test samples, up to this limit, by concentrating up to this limit and test for irreversible aggregation by diluting a concentrated sample to a number of levels and test for monodispersity. With the sensitivity of current DLS equipment even samples at 10 mg/ml should be measurable. In this way and within the limits of your DLS machine it should be possible to find out whether you will have aggregates when you dilute your concentrated protein 2-fold when setting up a hanging drop. 
Anil Mistry 

Pickled Crystals‹Unusual Additives! (A True Story) 
Pickle juice was added as an additive to a mutant form of a crystallized native protein. The mutant could not be crystallized in near similar conditions of the native. By chance, the components of pickle juice were read and found to contain compounds used in crystallization (i.e. Glycerol, PEG 400, citric acid, acetic acid, alum and a few vitamins). This juice (Sweet & Snappy Vlassic brand) was filtered (0.45 µl and pH¹ed to neutrality. It was then added to various PEG¹s (that crystallized the native form) and set up with the mutant. Crystals formed after 1 week! Trying to "add back" single components of the pickle juice to determine which component was responsible gave no crystals, the pickle juice (~1%) was necessary. Hint/ Tip: Commercially available food/ detergent solutions ought not to not be discounted as additives for crystallization! 
Michael Hickey 

Improve Your Crystals in Size, Shape, and Quantity 
After you have crystals open the coverslip, remove the mother liquid in the droplet, dissolve the crystal with 3 ul of H20 or buffer and add 3 ul of well solution. Close the coverslip. The crystal will appear again. My crystals diffract ~ 4 A, sometimes I get twins that diffract ~2.5 A. 
Nham Nguyen 

Oily drops 
Give yourself plenty of time to retrieve crystals from a hanging or sitting drop experiment. Add a drop of paraffin oil over the experiment drop. You will have plenty of time to mount crystals, test them with dye (remove a crystal to another drop of oil to dye so other crystals in the drop can still be used), crushed, etc. This works especially well when you find crystals that may be protein in an old dried out plate that cannot tolerate much additional evaporation. 
Joe Luft 

Sit and spin 
When doing microbatch screening, briefly centrifuge the plate at low speed. That not only ensures protein and precipitants mix and form a single drop, but also separates particulate matter. Any immediate precipitate is pelleted softly allowing the majority of the drop to be easier to visualize under the microscope. This might also generate two concentration phases ofprotein. Crystals may form from the precipitate or the clear phase. 
Dean Devonshire 

Crosslinking 
Original suggestion from Steve Swerdon, NIMR. For crosslinking crystals, put glutaraldehyde into the moatt of a Douglas Instruments VaporBatch plate to crosslink crystals in droplets dispensed under Al's oil. 
Lesley Haire 

Two to one 
In setting up nanoliter screens use 2:1 protein:reagent. Higher hit rate. Three examples so far. 
Heather Ringrose 

Smooth operator 
Smooth soaking in agarose gel. Grow your crystals under the usual conditions with 0.1 to 0.3% agarose gel and let the soaking solution (ligands, heavy atoms) diffuse smoothly through the gel. 
Claude Sauter 

Light em up 
Use luminal in protein crystal identification. Luminol is used in forensics to stain blood. Uses UV to light up blood stains. Add luminal to drops with crystals. Luminol binds to protein crystals. Addition of UV light and protein crystal will fluoresce. Salt crystals should not fluoresce. 
Christopher Browning 

What about it 
What about the production of protein crystals in transgenic yeast containing the genes of Bacillus thuringiensis. Bacteria too small. Yeaste cells of reasonable size. Bacillus thuringiensis produce protein crystals in vivo, thus utilize this technique to produce crystals in yeast with overexpression of target protein. Or oocytes. 
Christopher Browning 

Silicon carbide 
Seeding drops with silicon carbide. Silicon carbide whiskers used to deliver DNA into cell nucleus. Microscopic in size but contain suitable surface for protein molecules to aggregate and nucleate for crystal growth. Uniform size of whiskers will allow reproducible crystallization set ups. Thereby could be used in initial screening trials to induce nuclei for crystals to grow. 
Christopher Browning 

Seeding 
Use seeding for screening when you have only precipitate in your first screens. 
Jens-Christian Poulsen 

Don't change 
If you are having problems repeating or transferring crystallizations from 96 well to 24 well plates, for optimization stick with the 96 well plates. The single well low profile Greiner plate case easily be set up by hand or robot. There is plenty of room for larger drops and crystals are accessible for fishing. There is even a ledge for a small drop of cryo.
Shirley Roberts 

Glass seeds 
Seeding with pieces of glass, broken cover slides. The protein crystals sometimes grow along the end of the glass. 
Gaby 

Water replacement 
If cryo-cooling is not giving satisfactory results, check how you are making up the cryoprotectant solution. The water in the mother liquor should be replaced by the cryoprotectant agent, rather than diluting the mother liquor. This factor is the single most common factor causing trouble which is easily rectified. 
Elspeth Garman 

Be the crystal 
When manipulating crystals from their growing drop, and soaking them, try to imagine that you are the crystals, and how you would feel being poked with a loop or a needle, or what it would be like to have the pH temperature, osmotic pressure around you suddenly change without warning. It might help your procedures! 
Elspeth Garman 

He's not heavy, he's my crystal 
Wrap some lead tape (available from golf shops) around the lid of a cryo cap then the cap sits the appropriate way in a dewar of liquid nitrogen for easy storage of your frozen crystal. 
Janet Newman 

Fluidigm optimization 
When optimizing leads obtained from the Fluidigm chips in a vapor diffusion format, screen different ratios of protein to well volumes in drops. 
Neil Grodsky 

Microbatch 
If your protein is not very concentrated, set up microbatch screening experiments with 0.1 microliter screening solution plus 0.5 microltier protein. Use Al’s oil to allow concentration in the drop. That way you are less likely to get salt crystals before you get protein crystals. 
Patrick Shaw Stewart

No hits, what next 
No hits from a screen, what next? You can always set up another screen! Or, you can make a list of all drops that have precipitate and use the precipitate as a potential seed stock. Streak seed from the precipitate into new drops that have been set up at 50% and 75% of the original screening solution. There may be nucleation sites or microscopic seeds in the precipitates that may grow at lower precipitate saturation. Better still, streak seed from the precipitate to a clear zone into the same drop to recycle the drop. 
Anil Mistry 

Reproducible seeding 
For reproducible micro-seeding by hand use a cryoloop to fish out your seed from the seed stock and transfer them to the drop. Use a 0.3-0.4 mm cryoloop. 
Anna Aagaard 

Reduce to enlarge 
For bigger crystals try to add 0.5-1 microliter of 14 M beta-mercaptoethanol to the reservoir after the protein drop was set up. 
Zhanna Druzina 

Difficult ligands 
Problem: Can grow crystal but no protein-ligand crystals. Tip: Take the conditions from the apo crystals and develop a focused optimization screen (24 well maximum). Screen complexes using cross seeding and the focused screen and three drop ratios (1:1, 2:1 and 2:3). 
Annie Hassell 

Time for a change 
Problem: Poor crystal quality. Tip: Change tray type or crystallization method. For example, initial screens done in sitting drop tray and crystal quality improved in 96 well hanging drop tray. 
Annie Hassell 

Avoid the rut 
Every project, every protein, every construct is unique. Be careful of knowing too much. Just because things did or did not work in the past does not mean things will work that way for the next project. 
Brandon Collins 

Get heavy to stabilize 
Problem: Poor diffraction. Tip: Heavy atom soaks to stabilize floppy regions of the protein. 

Lose focus 
Don't focus all of your optimization efforts on a single crystallization condition. If you have several different crystallization conditions identified for a sample go after them. Crystals of the same protein produced from different chemical conditions and/or temperatures will have unique physical properties. These properties will determine how easy the crystal can be looped (physical stability), cryoprotected and ultimately how well the crystal diffracts X-rays. Avoid single points of failure, go after several hits. 
Joseph Luft 

Ligand seeding 
If ligands can't be soaked into crystal or co-crystallization is ligand specific try seeding into drops that contain ligand of interest. When soaking crystals with insoluble ligands try adding the cryoprotectant to the soaking solution. This can help solubilize the ligand and also cryoprotect (so less handling). Doesn't work so well when salt is the cryoprotectant, but may well when it's glycerol, DMSO or ethylene glycol. 
Doug Marcotte 

Mostly clear 
Problem: Drops are all/mostly clear. Tip: Remove stabilizing agents (salt, glycerol, etc) from the protein buffer. Then do crystallization screens. 
Annie Hassell 

Hydrate or die 
At suboptimal protein concentrations the interface between protein solution and crystallization screen solution may exhibit excessive precipitation. To avoid this, before adding screen solution add 1 microliter of water. Downside of this is equilibration will take slightly longer. Upside of this is decreased osmotic shock for protein and less precipitation. 
Vaheh Oganesyan 

What failure looks like 
Put one conditions of 40% TCA into your standard screen. This should precipitate out all of your protein, so that you have an idea of what heavy precipitate should look like. 
Janet Newman 

Think global act local 
Don't consider a crystallization result in isolation. Look for neighbors in chemical space and use those results to provide chemical directions for optimization. If a cryocooled crystal does not diffract well. You cannot tell if it is the crystal, cryoprotectant or cooling that is causing the problem. Look at room temperature data before moving on. 
Edward Snell 

X marks the spot 
Consider the case of poor nucleation and seeding did not work. Tip: Mix protein and well solution then use pipette tip to cross the drop into branched shape. Crystal may grow in the branches of the drop. 
Mei Xu 

Robotic seeding 
To increase you choices of producing more optimal crystal condition or conditions using seeds, try the following. Program a small volume liquid handler to dispense your protein and seed solution directly into 96 well commercial screens and/or an additive screen. 
Paris Ward 

Delete to succeed 
Recent success with loop deletions. Sequence alignments reveal either charged loops and/or loop insertions relative to homologues. I have removed 3 to 34 amino acids and retained high expression soluble protein and novel crystals.
Heidi Schubert 

PEG to the rescue 
Problem: Unstable protein. Tip: Add 1-5% low molecular weight (200 to 1,000) PEG directly to the protein and then screen. 
Annie Hassell 

Doh! 
Start with big crystals! Add at least 5% glycerol to everything. 
Jim Pflugrath 

Cryo stabilization 
Stabilize crystals in cryo by adding protein buffer components into the cryo. For example, most commonly I add 100 to 150 mM NaCl plus reservoir components plus cryoprotectant(s). 
Laura Pelletier 

Play with the ratio 
Play with protein-mother liquor ratio, especially with low solubility proteins. Try different concentration of protein coupled with streak seeding. 
Ayse Sinem Ozyurt 

Repeat after me 
After looking at the results of initial screening or of additive screen, pick several of the best "hits" and screen in 96 well format, 6 or 8 identical drops of each favorite before scaling up. Helps to eliminate "one offs" and save time. 
Nancy Bump 

Complex it 
Apo protein is monidisperse but won't crystallize. Complex it! Complex it! Complex it! 
Paul Reichert 

Matrix seeding 
If your crystal seeds withstand large serial dilutions, try matrix seeding via the reservoir by doing the following. 1) Create crystal seeds as described by Allan D'Arcy. 2) Dispense seed into reservoirs containing reservoir. 3) Aispirate/dispense to mix seed in reservoir. 4) Dispense mother liquor droplet containing seed onto protein drop. 
Armando Villasenor 

Salt that sucker out 
If you do not see any crystal growth in several days after set-up (more than 1 to 2 weeks) and the drop are not all clear, add salt (such as ammonium sulfate) to 0.5 M to the drops. Even though ammonium sulfate salt crystals might form, you might actually get protein crystals. This worked for me recently. 
Neil Grodsky 

Helpful urea 
Non denaturing (less than 3 M) concentrations of urea can be helpful to solubilize your protein. 
Gloria Borgstahl 

Precise dosing 
Try stoichiometric levels of multivalents, cations as additives. They may be necessary for crystallization but sometimes the levels found in commercial screens are too high and toxic. 
Simon Low 

Cook to crystallize 
Heat treatment of protein complex to obtain diffraction quality crystals. Original complex has no initial crystal hits. Heat treat protein complex (25 to 80 degrees Celsius) for various times (5 to 30 minutes). Centrifuge to get rid of aggregated protein and screen again. 
Liping Wang 

Gap it 
When using pre greased trays, take a toothpick and remove a bit of grease from each well. Now as you push down on your cover slip, you turn it a few degrees. This will allow the air to escape and the turn will form an airtight seal over each well. 
Paul Reinfelds 

Thermostability 
Thermostability. Monitor the effect of additives, buffers, ligands, etc. on melt temp of your protein. We have seen in multiple cases that the most thermostable construct, buffers, additive yields the best or only crystals. How? We use Bio-Rad's iQ5 iCycler (a PCR instrument) as it has 5 sets of filters for excitational emission and hydrophobic dyes that fluoresce upon binding (protein unfolding). 
Jackie Day 

Sticky situation 
Problem: Crystals adhering to plastic of sitting drop plate, and mechanical dislodging (by cryoloop, tool, etc) does not work. Solution: Stan a fine gauge syringe needle into the plastic, near but not into the crystal. This often distorts/disrupts the plastic near the crystal and breaks the seal. 
Michael Wiener 

Big low tech 
Getting bigger crystal by low tech / low cost counter diffusion. If you are faced with either no nucleation or showers of crystals and the usual tricks including seeding do not work, try this: On a cover slide, set the protein and precipitant drop (example 1 microliter plus 1 microliter) separate, but very close to each other. Then, with a whisker or pipette tip streak through the drops to form a connecting bridge between the protein and precipitant solution. Invert cover slide and place over well. Crystals will form along the gradient and "self screen" for best conditions. 
Margarete Neu 

Insoluble compounds 
When working with compounds for co-crystallization, if the compounds are highly insoluble in protein buffer (50 micromolar or less) we often employ low concentration complexing. We dilute the protein and then add in diluted compound, so tha the compound is added close, or at least closer to a concentration where it is soluble and the content of DMSO in the protein sample remains less than 2%. The protein-compound complex is then concentrated for crystallization trials. This has helped us with several projects with highly insoluble compounds. 
Elizabeth Fry 

Ionic liquids 
In experiments using model proteins we found that Ionic Liquids (IL's) specifically 1-Butyl-2-methyl imidazolium chloride gave increased numbers of crystallization outcomes compared to the IL controls. A large number of the crystals obtained had precipitated outcomes in the IL controls. In many other cases the IL and crystal had an improved morphology (needles to plates, plates to 3D crystals) over the IL controls. Tip: Using an IL such as 1-Butyl-2-methyl imidazolium chloride as an additive to improve chances of getting a crystal from conditions which otherwise would give precipitate. Marc Pusey, MI Research, Inc. Our favorite cryoprotectant. 1x UCP (Ultimate Cryo Protectant) 8% Glycerol, 8% Ethylene glycol, 9% Sucrose, 2% Glucose. We make a 2x solution. Generally add this 1:1 with reservoir. The ratio can be modified, for example 1.2 microliter 2x UCP : 0.8 microliter reservoir, or 1.4 microliter 2x UCP : 0.6 microliter reservoir, or 0.6 microliter 2x UCP : 1.4 microliter reservoir, etc. I believe this has been successful in cryoprotecting some 70% of all of our systems, resulting in more than 50 solutions of these targets regardless of previous cryogenic treatments. Author in unknown to me, but the credit is published in a singled Hencrickson paper. I was tipped off 6 years ago. 
Christopher Bonagura 

Se-met stuff 
If you can't get your Se-met protein to crystallize try leaving ou the DTT/TCEP during purification and crystallization.Sometimes there are disulfide bridges near crystallization contacts that need preserving for crystallization to take place. Collected the Se edge is still possible. 
Shirley Robert 

DMSO cryo 
Try 20 to 30% DMSO as cryo. This has worked well in a number of cases for me and I've added this to a very short list of cryos that I personally use. Note: If your mother solution contains a high salt concentration the DMSO will cause it to precipitate out of solution. So beware! 
Rich Romero 

Stack em up 
4 degrees Celsius crystallization plates prepared at room temperature always have a condensation problem on the plate seal. To avoid condensation cover the finished plates with two lids and plate it in the cold room for 20 minutes or on top of a cold metal block, The will reduce the temperature of the reservoir solution while the 2 lids delay the temperature change from the top long enough to avoid condensation. 
Beat Blattmann 

Anti skid 
Ever been manually sealing a plate only have it slip out from under you? The result is usually death for hanging drop plates, and with sitting drop plates your best bet is hoping the drop is not splashed onto the seal above. To ensure the plate stays put when applying pressure, we use a "grip pad" in our lab. Simply place the grip pad onto the bench, set plate on top and seal as usual. The grip pad prevents the plate from slipping out from under the compression tool, usually a brayer, used for ensuring the please seal is applied correctly. The grip pad can be cut from the material commercially available for lining tool shop drawers. In addition, we created a fixed plate holder that encloses the entire plate to guarantee the brayer does not slip off the plate when sealing, a common occurrence when manually sealing many plates. Our plate holder is custom cut from a hard rubber to fit both thee 24 well and 96 well plates. The plates sit slightly above the platform to ensure both ends are sealed and for easy removal. The sides of the platform are rounded to ensure the brayer has a smooth path of travel. 
Barbra Pagarigan

Should one remove 1 M imidazole from the sample after elution from a nickel column? In many cases it is good to have (an) additional purification step(s) following the elution of the sample from a Ni column. Size exclusion chromatography (SEC) is a convenient and effective way to remove the imidazole. If one finds that imidazole is an important variable in maintaining sample homogeneity, one can also simply reduce the concentration of imidazole from 1 M to a more reasonable level (0.02 – 0.2 M) for inclusion of crystallization reagents. In some cases a protein might not crystallize in the presence of imidazole. To remove the imidazole one can try precipitating the protein in ammonium sulfate and resuspending the sample in low salt buffer and purify by ion exchange or SEC. One can also remove imidazole by dialysis. In one instance (personal communication from Artem G. Evdokimov) decent crystals could only be obtained using 0.6 M imidazole/acetate buffer and two imidazole molecules were seen occupying hydrophobic spots on the protein surface. Sometimes the reason Ni column purified proteins precipitate without imidazole is that the Ni ions leak from the Ni column. The presence of imidazole may have been chelating this excess and trace Ni and once removed the remaining Ni Ni may cause the protein with a His tag to aggregate in solution. One may be able to remove this excess Ni using a Ni chelating resin (Hampton Research catalog number HR2-312). Or one can leave the imidazole buffer with the sample in a more reasonable concentration (0.02 – 0.1 M) or try citrate buffer (chelator) or EDTA. If you are working with a metalloprotein and need metals in the sample, using the chelating resin rather than leaving a chelating reagent in the sample is obviously a better choice.

If a protein does not crystallize or gives poorly diffracting crystals one might try heat treatment of the sample. This is a quick experiment with minimal equipment requirement. The procedure can reduce and/or eliminate protein that is not folded properly from the sample. Place the sample at 4 degrees Celsius for one hour. Incubate the sample at 37 degrees Celsius for one hour. Centrifuge the sample or filter using a 0.22 micron filter. Screen the heat treated sample for crystallization. The time course and temperature may require optimization for nest results. One can monitor the heat treatment using dynamic light scattering or an activity assay. Tip from Annie Hassell, GSK – IUCr 2005.

The "solution" we hit upon for a problem protein with a high solubility profile was to cocrystallize it with an Fab fragment. Not the easiest route, but it worked. Catherine L. Lawson, Rutgers University. Reference: Li H, Lawson CL. (1995) Crystallization and preliminary X-ray analysis of Borrelia burgdorferi outer surface protein A (OspA) complexed with a murine monoclonal antibody Fab fragment. J Struct Biol. (115 )335-337.

Compounds produced in medicinal chemistry efforts often have low aqueous solubility when compared to biological ligands. When poor compound solubility is suspected, one may try dimethylsulfoxide (DMSO) to solubilize the compound. Dissolve the compound in 100% DMSO and THEN dilute to a lower DMSO concentration. Dissolving a compound in dilute DMSO can be a very slow kinetic process. 

One may consider evaluating other solvents for compound solubility, including Polyethylene glycol 200, ethylene glycol, 2-propanol, or methanol. 

Another option is to evaluate adding the dry compound to the drop as has been done successfully with numerous insoluble heavy atom derivatives. Although the compound may not appear to be soluble in the drop, there may be some solubility, sufficient for the compound to bind the protein.

Add 50 mM L-Arginine together with 50 mM L-Glutamic acid to the protein sample to improve protein solubility and long-term stability. Golovanov et al reported a 3 to 8 fold improvement in solubilization with six different test proteins. 

Reference: A simple method for improving protein solubility and long-term stability. Alexander P. Golovanov, Guillaume Haubergue, Stuart A. Wilson, and Lu-Yun Lian. J. Am. Chem. SOc. 2004, 126, 8933-8939.

For most soluble proteins, 10 to 15 mg/ml in a sample buffer that promotes the sample stability, homogeneity and monodispersity will work fine for an initial crystallization screen. 

A pre crystallization test such as the Hampton Research PCT (catalog number HR2-140) can be used to better determine the appropriate protein concentration for crystallization screening.

The minimal number of chemicals and the minimal concentration of these chemicals necessary to promote sample stability, homogenity and monodispersity. 

Use a buffer concentration between 10 -25 mM. This will allow crystallization reagents with a buffer concentration of 50 - 100 mM to manipulate the pH of the sample during the experiment. 

Current trends seems to favor the use of 200 mM or less NaCl or KCl in the sample. Using no NaCl or KCl is just fine as well. It simply seems most overexpressed proteins are purified and prepared into a final sample buffer contain NaCl. 

If the sample is known or expected to have free sulfhydyl residues on the surface, including a reducing agent such as TCEP hydrochloride can help to prevent oxidation of these residues. Oxidation of sulfhydryl residues can lead to sample aggregation. Avoid reducing agents if the sample has one more more disulfide bonds in order to prevent cleaving these disulfide bonds. 

If your screens contain di or poly valent cations such as calcium, zinc, iron and others, avoid phosphate, borate and carbonate buffers. Such mixtures can results in false positive salt crystals.

The crystallization of proteins with phosphorylation sites can often be difficult. It is imagined that such samples are susceptible to heterogeneity due to the presence of multiple species of protein in various states of phosphorylation. Careful sample purification and preparation is perhaps more of a significant crystallization variable rather than any particular crystallization trick or specialized screen. One can use a strong anion exchange chromatography with a shallow elution gradient to prepare a homogenous sample for crystallization. Isolation of a single phosphorylated form can produce crystals where as the presence of multiple phosphorylated forms can lead to precipitate, microcrystals or crystals less suitable for diffraction compared to those from a homogeneous sample. One may use mass spec to characterize the different elution samples following chromatography to assist in the selection of the best sample for crystallization.

Do not mix different batches of protein. Each batch of protein should be considered and treated separately. The reason being that the expression and purification conditions as well as the chemical and physical conditions are not identical for each batch. Even subtle differences between each batch can affect the outcome of a crystallization experiment. Each batch should be characterized and documented. Run at least an SDS-PAGE. If possible, consider also a native PAGE, IEF-PAGE, analytical gel filtration, mass spec and dynamic light scattering to characterize each batch. Save some protein from each batch for later experimentation, comparison, documentation and reproducibility.

Selenomethionyl proteins can be more sensitive to oxidation than natural proteins. If selenium atoms are on the surface of the protein molecule they can alter protein solubility and hydrophobicity (typically less soluble and more hydrophobic). Such changes can affect the solubility and crystallization of the protein. 

To avoid oxidation of selenomethionine, buffers can be degassed. Buffers can also include a reducing reagent such as dithiothreitol (DTT) or beta-mercaptoethanol (BME) or TCEP hydrochloride as well as a chelator such as Ethylene Diamine Tetraacetic Acid (EDTA) to remove traces of metals that could catalyse oxidation. 

To slow oxidation of the sample, store the protein in a reducing environment of 1 to 10 mM DTT or beta-mercaptoethanol (BME) or TCEP hydrochloride. One can also add excess methionine towards preventing oxidation of Se-Met.

Proteins are sensitive to oxidative damage, often with important biological effects as well as affects on the protein's ability to crystallize. 

Methionine, cysteine, tryptophan, tyrosine and histidine residues are susceptible to oxidation. 

The oxidation of susceptible resisude presumably affects protein structure and stability by reducing side chain hydrophobicity, increasing the capacity for hydrogen bonding and altering the size and shape of the amino acid. 

Oxidation can be slowed and minized by including reducing agents in the sample buffer and the crystallization reagent. Appropriate reducing agents include dithiothreitol (DTT), beta-mercaptoethanol (BME) and Tri(2-carboxyethyl)phosphine hydro-chloride (TCEP hydrochloride). Free methionine can be employed as an effective antioxidant for samples containing methionine. Typical reducing agent concentration is 1 to 10 mM. 

Oxidation prone residues can also be removed and replaced by mutagensis towards engineering oxidative resistance in the protein.

Say your protein is eluting from the column (IMAC - Immobilized Metal Affinity Chromatography) as an aggregate and showing more absorbance at A260 than A280 or a lower than desired 280/260 ratio, indicating the presence of nucleic acid. Unless you’re working on a protein nucleic acid complex, this nucleic acid could be considered a contaminant that might be the cause of the aggregation of your sample or the contamination would interfere with the crystallization of the sample. To remove the contaminating nucleic acids, add DNase to the sample prior to loading onto the column and increase the column wash to see if this can remove the contaminating nucleic acid, resulting in an increased 280/260 ratio. 

Avoid repeated freeze thaw cycles with your sample. 

Freezing is best done quickly. Snap freeze small aliquots (50 to 100 microliters) in thin walled PCR tubes. (An improved protocol for rapid freezing of protein samples for long-term storage. Deng J, Davies DR, Wisedchaisri G, Wu M, Hol WG, Mehlin C. Acta Crystallogr D Biol Crystallogr. 2004 Jan;60(Pt 1):203-4. Epub 2003 Dec 18.) 

Alternatively, one can use a 200 microliter pipette set to the desired volume (30 ul for example) and pipette drops of protein directly into a small Dewar of liquid nitrogen (for example, fill 100 ml of liquid nitrogen into a 500 ml container). Avoid plastic as some proteins will adhere to plastic when frozen. Use safety glasses. The pipetted protein will form small frozen pellets. Pipette slowly, allowing the drops to freeze solid before adding the next one. The liquid nitrogen can be poured off, allowed to boil or or the frozen pellets can be picked up from the liquid nitrogen with forceps and stored in a cryovial in the –80 freezer. In this format, one can thaw only the amount of sample needed. 

Thawing is best done quickly. Hold the PCR tube under running cold water after removal from the freezer and then place the sample in the ice bucket after it has completely thawed. 

It is a good idea to check sample homogeneity, stability and activity after thawing, especially after long term storage.

Biochim Biophys Acta. 2006 Sep;1760(9):1304-13. 
Structural analysis and classification of native proteins from E. coli commonly co-purified by immobilised metal affinity chromatography. 
Bolanos-Garcia VM, Davies OR. 

Ferric uptake regulator (Fur) 
Metal-binding lipocalin (YodA) 
Cu/Zn-superoxide dismutase (Cu/Zn-SODM) 
Acetylornithinase (ArgE) 
Glycogen synthase (GlgA) 
Carbonic anhydrase (YadF) 
Glucosamine-6-phosphate synthase (GlmS) 
cAMP-regulatory protein 
(CRP) 
Host factor-I protein (Hfq) 
Chloramphenicol-O-acetyl transferase (CAT) Peptidoylproline cis–trans isomerase (SlyD) Regulatory ribosomal protein (S15) Formyl transferase (YfbG) Glucose-6-phosphate 1-dehydrogenase (G6PD) GroEL/Hsp60 Component 1 of the 2-oxoglutarate dehydrogenase complex (ODO1) Component E2 of the dihydrolipoamide succinyltransferase (ODO2) 
Glucose-6-phosphate 1-dehydrogenase (G6PD) 
Glucose-6-phosphate 1-dehydrogenase (G6PD) 
inorganic pyrophosphatase 
lac repressor 

Acta Crystallogr Sect F Struct Biol Cryst Commun. 2007 Jun 1;63(Pt 6):457-61. Epub 2007 May 5. 
Purification, crystallization and structure determination of native GroEL from Escherichia coli lacking bound potassium ions. 
Kiser PD, Lodowski DT, Palczewski K. 

Protein Expr Purif. 2010 Apr;70(2):191-5. Epub 2009 Nov 1. 
Identification and characterization of native proteins of Escherichia coli BL-21 that display affinity towards Immobilized Metal Affinity Chromatography and Hydrophobic Interaction Chromatography Matrices. 
Tiwari N, Woods L, Haley R, Kight A, Goforth R, Clark K, Ataai M, Henry R, Beitle R. 

For membrane proteins purified from E. coli AcrB can be a problem, as well as ferritins, Omp porins and succinate dehydrogenase. 

J Struct Biol. 2009 Apr;166(1):107-11. 
AcrB et al.: Obstinate contaminants in a picogram scale. One more bottleneck in the membrane protein structure pipeline. 

Acta Crystallogr Sect F Struct Biol Cryst Commun. 2008 Oct 1;64(Pt 10):880-5. 
There is a baby in the bath water: AcrB contamination is a major problem in membrane-protein crystallization. 

Biochim Biophys Acta. 2006 Sep;1760(9):1304-13. 
Structural analysis and classification of native proteins from E. coli commonly co-purified by immobilised metal affinity chromatography. 
Bolanos-Garcia VM, Davies OR. 

Acta Crystallogr Sect F Struct Biol Cryst Commun. 2008 Oct 1;64(Pt 10):880-5. 
There is a baby in the bath water: AcrB contamination is a major problem in membrane-protein crystallization. Veesler D, Blangy S, Cambillau C, Sciara G.

Solubility suggestions for reconstituting a lyophilized peptide. 

Use sterile water. 

If there are any Methionine (M), Cysteine (C), or Tryptophan (W) residues, use oxygen free solvents to prevent oxidation. 

Salt can hinder peptide solubility. Avoid phosphate buffered saline (PBS). 

For peptides with hydrophilic residues (KRHDEN; Lysine, Arginine, Histidine, Aspartic acid, Glutamic acid, Asparagine) try water. 

For peptides with hydrophobic residues (AVLIMFW; Alanine, Valine, Lysine, Isoleucine, Methionine, Phenylalanine, Tryptophan) try DMF, DMSO, TFA or Acetonitrile since hydrophobic residues have low solubility in aqueous solvents. 

If a peptide with hydrophilic residues is not soluble: 
1) Adjust the pH according to the overall charge of the peptide. 
a) Count the possible positive charges (KRH; lysine, arginine, histidine, and the free N-terminus) 
b) Count the possible negative charges (D,E; aspartic acid, glutamic acid, and the free C-terminus) 
c) Determine which is greater 
e) If positive charges are greater, add dilute acid to protonate residues and maximize charge. If negative charges are greater, add dilute base to deprotonate and maximize charge. 
2) Gently warm the sample 
3) Sonicate the sample 
4) Add an organic solvent such as DMSO, Acetonitrile, DMF or ethanol.

If the peptide in insoluble in water, dissolve the peptide in DMSO, up to 100 mg/ml. Prepare a 10:1 peptide:protein solution with the DMSO reduced to 2% or less. Centrifuge and observe for precipitate to gauge the solubility of the peptide protein complex. Set crystallization screens with the peptide protein complex.

Evaluate how different protein concentrations influences the precipitation of the sample after storage. Generally it is better to store proteins more concentrated than dilute. Losses due to adsorption will be relatively less when the sample is at 10 mg/ml than at 0.5 mg/ml. However, if storing the protein too concentrated leads to precipitation, store the protein at a dilute concentration and then concentrate the sample immediately after thawing and right before crystallization experiments. 

In some instances a protein will crystallize during a freeze thaw so be sure to check the precipitate under magnification for the presence of microcrystals. 

Try warming a precipitate protein by holding or rubbing the tube in your warm hands or holding the tube under a stream of warm (approximately 37 degrees Celsius) tap water with gentle swirling. Be gentle. Do not shake or vortex. Avoid foaming.

Try to crystallize the glycosylated and deglycosylated protein. Some proteins do crystallize with full N-linked glycans. 
If one does not have enough protein to try everything, or if the glycosylated protein will not crystallize, deglycosylation should be tested. There are at least two things to consider when deglycosylating a protein. First, how easy or difficult it will be to deglycosylate the protein to homogeneity and second how much less soluble the protein may become after glycosidase treatment. 

Try different glycosidases. Try PNGase first as it removes the whole N-linked glycan. Then try the endoglycosidases such as Endo F1, F3, Endo H, which leave a single GlcNAc on the Asn. If the endoglycosidases do not work, then consider exoglycosidases such as sialidase, galactosidase, hexaminidase, and mannosidase. In general, the smaller the resulting N-glycan residue, the less flexibility you will have left on the protein, meaning a higher degree of conformational homogeneity and a likely higher chance of getting crystals (that diffract). 

The chemical and physical conditions one will need for an efficient cleavage with any certain enzyme can vary. This is most likely due to the accessibility of the cleavage site. In the most facile cases, treatment at 4 degrees Celsius with trace amount (1:10,000) of PNGase O/N can result in complete removal of the N-glycans. On the other extreme, for certain proteins one may need more enzyme than substrate protein and the digestion performed at 37 degrees Celsius. Varying incubation time, enzyme concentration, sample concentration and temperature can be used to achieve full deglycosylation. 

Being the enzyme that cuts the deepest on the N-glycan, one problem with PNGase is that the cleavage site (the NH2-C linkage on the Asn side chain) on certain N-linked glycosylation site could be quite inaccessible. In many cases, one may find one has to heat the reaction to 37 degrees Celsius in order to achieve a certain level of PNGase cleavage. For highly N-glycosylated proteins (having more than 4 or 5 sites), PNGase treatment is almost sure to fail. The other thing is, PNGase treatment often results in poorly soluble proteins as the removal of the N-glycan exposes some surface patch for aggregation, or destabilizes the protein. 
The Endo F1, F3, H, especially Endo H, can often result in better completeness of N-glycan removal, and better solubility of the protein, with the expense of one GlcNAc left on the protein. However, these can require the N-glycans to be of certain type. For example, Endo H requires the N-glycans to be high-mannose type, which has to be generated from certain cell lines, or by a series of exo-glycosidase treatment. Note: For insect cell produced proteins, endo H is very likely to work. 

One can often achieve some degree of separation of the glycosylated form and un-glycosylated on ion-exchange, even when the N-glycans are not the charged type (for example, high mannose). This is likely due to a solubility difference of the differently glycosylated species. 

In cases where one has charged N-glycans, for example, proteins made from normal mammalian cells, the negative charge of the sialic acid on the N-glycans could be significant, that it can mask the protein's own surface charge. 
Consequently, the protein, regardless of its real PI, will bind to ion exchange resin, while the deglycosylated will still behave normally. This can result in the separation of the two or more species. Actually, highly glycosylated proteins with complex type glycans (with terminal sialic acid) often run as multiple peaks on ion exchange, due to the heterogeneity of the N-glycan. 

Hydrophobic Interaction Chromatography (HIC) should also be considered as a way to separate the deglycosylated protein. 

Lectin columns are another consideration when is comes to separating the deglycosylated protein. However the applicability of lectins depends on the N-glycan's type, and this can get complicated. 
Proteins produced from insect cells are often high-mannose type. This means that the N-glycans from insect cells are likely susceptible to Endo H treatment, that the N-glycans are not charged, that the N-glycans themselves are more or less homogenous as they are likely to be Man3,4,5 only, and are also smaller than the complex type glycans made from mammalian cells. This would lead one to believe that such glycosylated proteins are more amenable to crystallization. 

Keep in mind that 100% purity and homogeneity, although ideal is often not required for crystallization. Keep in mind that moderation is key and that one might be better off with more partially purified sample screening than less ultrapure sample for screening. 

When starting our screening glycosylated and deglycosylated proteins it is generally a good idea to try a chemically diverse screen such as the Hampton Research Index screen. Based upon the results of Index, one may then proceed to the appropriate chemical space identified by Index, meaning salt based, polymer based, mixed polymer/salt, and so on. 

References 
Protein Glycosylation, Sweet to Crystal Growth? Jeroen R. Mesters and Rolf Hilgenfeld. Crystal Growth & Design, Vol. 7, No. 11, 2007. 

Strategies in the crystallization of glycoproteins and protein complexes. Stura, Enrico A.; Nemerow, Glen R.; Wilson, Ian A. Journal of Crystal Growth, Volume 122, Issue 1-4, p. 273-285 (1992) DOI: 10.1016/0022-0248(92)90256-I 

An efficient platform for screening expression and crystallization of glycoproteins produced in human cells. J. E. Lee et al. Nature Protoc. 4, 592-604 (2009). doi:10.1038/nprot.2009.29.

To convert mg/ml to mM divide the concentration in mg/ml by the molecular weight of the sample and multiply by 1,000. 

For example, a 10 mg/ml solution of lysozyme is 0.68 mM 

Approximate molecular weight is 14,700 g/mole) 

10 mg/ml x 1 g/1,000 mg x 1 mole/14,700 gram x 1,000 ml/1 L x 1,000 millimole/1 mole = 0.68 millimolw/ml = 0.68 mM.

Applications 
Crystallization grade NDSB-195 for formulating screens or for optimization 

Features 
Efficiently solubilizes proteins at non-denaturing concentrations 

Useful in preventing protein aggregation 

Non detergent sulfobetaines (NDSB) are used for protein folding, renaturation and crystallization 

Description 
The NDSB are a group of zwitterionic compounds that can reduce aggregation and aid in refolding proteins found in inclusion bodies and bacterial expression systems. Non detergent sulfobetaines have a sulfobetaine hydrophilic group and a short hydrophobic group that cannot aggregate to form micelles, therefore NDSB 's are not considered detergents. NDSB increase the extraction yield ( up to 30%) of membrane, nuclear and cyto-skeletal associated proteins. The short hydrophobic groups combined with the charge neutralization of the sulfobetaine group results in higher yields of membrane proteins. NDSB have been used in refolding and renaturation of chemically and thermally denatured proteins. 

The non detergent sulfobetaines are zwitterionic over a wide pH range, easily removed by dialysis and do not absorb significantly in the near UV range. 

Typical useful NDSB concentration in protein sample is 0.5-1.0 M. 


References 
1. Goldberg, M.E., et al. 1995. Folding & Design 1, 21. 
2. Vuillard, L., et al. 1995. Anal. Biochem. 230, 290. 
3. Vuillard, L., et al. 1995. Biochem. J. 305, 337. 
4. Vuillard, L., et al. 1994. FEBS Lett. 353, 294. 

Hampton Research NDSB's 
HR2-703 NDSB-195, 5 gram 
HR2-701 NDSB-201, 5 gram 
HR2-793 NDSB-211, 5 gram 
HR2-791 NDSB-221, 5 gram 
HR2-705 NDSB-256, 5 gram 
NDSB-195 and NDSB-201 are also components of the Hampton Research Additive Screen (catalog number HR2-138 and HR2-428). 

Differential scanning calorimetry can give an absolute measure of protein thermostability. Thermofluor can assess the relative thermostability of a protein in different solvent conditions. 

Using Thermofluor, one may find that if the protein melts below 45 degrees Celsius that crystallization may be difficult or unsuccessful. Or it could indicate that one may set the crystallization experiments at temperatures below room temperature, such as 4 degrees Celsius. That being said, a protein melting above 45 degrees Celsius does not guarantee crystallization success. There are many other variables to consider in crystallization besides protein melting temperatures. 

Thermofluor can be used to screen pH, buffer type and solubilizing agents when searching for reagent conditions that promote sample stability. Protein stability can be an important crystallization variable and Thermofluor can allow one to rapidly screen many different pH levels, buffer types and solubilizing agents towards optimizing a sample buffer that promotes optimal sample solubility. 

In addition to the melting temperature one should also consider the shape of the melting curve. A very high initial flourescence can indicate a partially unfolded protein or hydrophobic patches on the surface of the protein. A very low initial flourescence can be a good sign compact proteins. A steep transmission curve is often considered better than a shallow surve. 

As a fluorescence reporter may likely interact differently with exposed hydrophobic patches in different proteins, one might be a bit more careful in comparing the Thermofluor results between different proteins. Yet, it can be a tool for reasonable comparison between proteins.

Suggestion 1 
Typically between 5-10x molar concentration over the protein is enough to ensure binding when the IC50 is uM to low mM. For tighter binding compounds (nM to low uM), 2-5x is sufficient. Whatever you do, when the precipitate occurs DO NOT REMOVE it. I learned to my chagrin that you change all the dynamics of the drop when you do. I ended up with empty crystals until I left the precipitate in place. Think of it this way: 

Free protein + compound ? protein:compound complex + precipitate (mix of protein + compound) 

If you change the equilibrium by removing the precipitate, you remove the “pressure” on the P:C complex, and it will dissociate to P + C. The precipitate acts as a reserve of protein and compound, thus favoring (or stabilizing) the P:C complex. I set drops up as a slurry frequently, and if I get crystals, they always have the compound bound. Pay attention to the drops if you are screening, because it will be important to note what makes the precipitated solution better (clear drops=solubilizing) or worse (aggregated drops=decreased solubility of your complex). You can also try suspending your compound in LMW PEG’s (200-400 FW) instead of DMSO. Either way, try using DMSO (~20%) or LMW PEG (~30%) depending on your crystallization conditions as a cryoprotectant. Any crystals that grow have some small amount of those agents in them already, so they should be more tolerant of them in higher concentrations. 

Reference: Bryan Prince, CCP4BB August 2011 

Suggestion 2 
If you do the calculations, you will find that you need a FREE ligand concentration of >10 * Kd to get >90% occupancy of the binding site. If you have e.g. a ligand with a Kd of 100 nM, you would need a free ligand concentration of 1 µM. However, a solution of 10 mg/ml of a protein of 30 kDa, has a protein concentration of 333 µM, so in theory you should have a total ligand concentration (free + bound) of 334 µM. In pratice, some of the ligand may have been degraded during (prolonged) storaged, or the compound may not be as pure as the chemist would have wished it to be, so it is wise to use a safety margin of at least two. We normally use 1-2 mM compound to be on the safe side. Having too much ligand usually does not hurt, except that you use more 

Reference: Herman Schreuder CCP4BB August 2011.

Add TCEP hydrochloride as a reducing agent to the sample buffer. TCEP hydrochloride does not interfere with IMAC columns and can help prevent multimerization through inappropriate disufide formation. 

Add 5 to 20% v/v glycerol during purification. 

Remove imidazole as soon as possible after nickel exchange chromatography. A number of proteins aggregate in the presence of imidazole, where others do not. 

Evaluate different buffer types as well as different pH conditions during purification. 

Evaluate different levels of ionic strength using sodium chloride. 

Evaluate adding lipids, detergents, substrates, inhibitors, ligands, co-factors, metal ions and other small molecule additives to promote monodispersity and minimize heterogeneity. 

If you are working with a membrane protein and basing the presence of heterogeneity on SDS-PAGE results, try not heating the sample before SDS-PAGE. Membrane proteins can aggregate in SDS-PAGE sample buffer, particularly when heated. This sometimes demonstrates itself on the SDS-PAGE as a dimer, ladder or smear. 

The heterogenity could be due to heterologous overexpression of a recombinant protein. Overexpression of a recombinant protein can lead to misfolded protein mixed with properly folded protein and heterogeneity. If one is overloading the membrane protein insertion system, one could reduce the induction.

MES, BES, CHES and PIPES buffers which possess a sulfonylethyl group, can contain a contaminant, oligo (vinylsulfonic acid), that is a side product in the chemical synthesis of these buffers. This polyanionic contaminant mimics RNA and at low salt (0.05 M Sodium chloride (NaCl) binds to and inhibits the enzyme with a Ki of 11 pM. This has implications to anyone studying nucleic acid-binding proteins. 

Reference 
Smith, B. D., Soellner, M. B., and Raines, R. T. (2003). Potent inhibition of ribonuclease A 
by oligo(vinylsulfonic acid). J. Biol. Chem. 278, 20934–20938. 

Try using surface plasmon resonance (SPR) to study the interaction between the protein and DNA. Use SPR to guide the choice of DNA for co-crystallization (Investigation of DNA sequence recognition by a streptomycete MarR family transcriptional regulator through surface plasmon resonance and X-ray crystallography. Stevenson C. E., Assaad A., Chandra G., Le T. B., Greive S. J., Bibb M. J., Lawson D. M. (2013) Nucleic Acids Research 41 7009-7022). 

If making and using different protein constructs, use SPR and Electrophoretic mobility shift assay (EMSA) to study whether the different protein constructs bind to the DNA. 

The length of DNA utilized in the crystallization experiments can be an important variable. Test a range of different lengths of DNA. Screen several different lengths of oligonucleotide with both blunt and sticky ends. Use SPR and try the shortest oligonucleotide that demonstrates good binding affinity. 

Desalted oligonucleotides are okay. Try them before spending more money on purified oligonucleotides. 

Try a slight excess of oligonucleotide to protein in the crystallization experiments. For example, Protein:Oligo 1:3. 

Purify the complex using size exclusion chromatography. Concentrate and screen the complex versus Hampton Research Natrix and Natrix 2. 

The BCA Assay works well with membrane proteins in the presence of detergents. 

The BCA assay is unaffected by typical concentrations of most ionic and nonionic detergents 

The BCA Assay is a detergent-compatible assay set to measure (A562nm) total protein concentration compared to a protein standard. The BCA Assay can be used to assess yields in whole cell lysates and affinity-column fractions, and samples ready for crystallization. Compared to most dye-binding methods, the BCA Assay is affected much less by protein compositional differences, providing greater protein-to-protein uniformity. 

The BCA Protein Assay combines the reduction of Cu2+ to Cu1+ by protein in an alkaline medium with colorimetric detection of the cuprous cation (Cu1+) by bicinchoninic acid. The first step is the chelation of copper with protein in an alkaline environment to form a light blue complex. In this reaction, known as the biuret reaction, peptides containing three or more amino acid residues form a colored chelate complex with cupric ions in an alkaline environment containing sodium potassium tartrate. 

In the second step of the color development reaction, bicinchoninic acid (BCA) reacts with the reduced (cuprous) cation that was formed in step one. The intense purple-colored reaction product results from the chelation of two molecules of BCA with one cuprous ion. The BCA/copper complex is water-soluble and exhibits a strong linear absorbance at 562 nm with increasing protein concentrations. The BCA reagent is approximately 100 times more sensitive (lower limit of detection) than the pale blue color of the first reaction. 

The reaction that leads to BCA color formation is strongly influenced by four amino acid residues (cysteine or cystine, tyrosine, and tryptophan) in the amino acid sequence of the protein. However, unlike the Coomassie dye-binding methods, the universal peptide backbone also contributes to color formation, helping to minimize variability caused by protein compositional differences. 

The original BCA assay is not compatible with reducing agents or chelating agents. There are now BCA assays that are compatible with reducing agents, although chelating agents are still incompatible. 

References 
Smith, P.K., et al. (1985). Anal. Biochem. 150: 76-85.

Dynamic light scattering, also known as DLS, also known as photon correlation spectroscopy, also known as quasi-elastic light scattering can be utilized to screen crystallization candidates for monodispersity. 

DLS measures the translational diffusion coefficient of a macromolecular undergoing Brownian motion in a solution. What? In essence, DLS measures the intensity of light scattered by molecules in solution. In turn, this measurement can tell you the size distribution of the protein molecules in solution. 

Crystal growers should use DLS to differentiate monodisperse solution from polydisperse solutions. Homogeneous, non-aggregated monodisperse proteins have a high probability of producing crystals. Proteins with non-specific aggregates and heterogeneous samples are less likely to crystallize. DLS allows one to screen protein samples quickly, with small volumes of sample, at different temperatures. The procedure is non-invasive so the protein can be recovered. If DLS is not an option one can obtain similar results using native polyacrylamide gel electrophoresis or size exclusion chromatography. But DLS is faster and more sensitive. 

For a great story on DLS as well as practical information on methods for DLS read Terese Bergor's chapter on Dynamic Light Scattering in Protein Crystallization - Techniques, Strategies, and Tips - A Laboratory Manual.

Avoid lyophilization of your protein for storage and concentration if you plan to use the protein for crystallization experiments. Lyophilization is a not so gentle procedure and can prevent crystallization. Yes, we know that lyophilized lysozyme will crystallize quite readily as well as other "Sigma proteins" but we have seen, as well as others, many a protein that will not crystallize following lyophilization. And we have yet to see a protein that would only crystallized after lyophilization. So don't go there. 

If you have a lyophilized protein there is hope. First of all, one should solubilize the protein in water or a stabilization buffer and dialyze the protein exhaustively against water or the stabilization buffer. Dialysis is an important step which will help to remove residual, non-volatile buffers and reagents as well as low molecular contaminants.

Mass spectrometers (MS) use the difference in mass-to-charge ratio (m/e) of ionized atoms or molecules to separate them from each other. MS is therefore useful for quantitation of atoms or molecules and also for determining chemical and structural information about molecules. Molecules have distinctive fragmentation patterns that provide structural information to identify structural components. 

MS can be a nice tool along with Dynamic Light Scattering (DLS), sodium dodceyl sulfate polyacrylamide gel electrophoresis (SDS PAGE), iso-electric focusing (IEF), and size exclusion chromatography (SEC) to evaluate the purity, homogeneity, and monodispersity of the sample prior to crystallization. Likewise, if crystallization problems exist, MS can be a nice tool to identify possible sources of the problem. MS can also be used to detect and possibly identify impurities in crystallization reagents. 

In the case of recombinant protein samples, an accurate measurement of the molecular weight will inform one about the presence of post-translational modifications. Dependent on the physical properties of the compound and the MS technique used, the molecular weight may be determined within an accuracy of one Dalton. The use of MS measurements will give sequence information where classical methods fail or may lead to ambiguous results as in the case of N-terminal blocked peptides, glycosylation or identification of the C-terminus. Next to this, MS is the ideal analytical method to support sample clean-up procedures and as a routine check for sample impurities of all kinds. As for sample requirements for MS, the samples should be free of non-volatile salts and the procedure typically requires between 1 ug and several nmoles, depending on the kind of analysis performed.

Try crystallizing your sample with a substrate, coenzyme, inhibitor, or ligand. Sometimes these agents serve to fix the macromolecule in a more compact and stable form. This may impose a greater degree of structural homogeneity and increase the likelihood of crystallization.

If you cannot get diffraction quality crystals of your protein, try both carboypeptidase-A and carboxypeptidase-B to trim the carboxy terminus of the protein in order to generate smaller protein fragments which might be more amicable to crystallization.

Use sequence grade trypsin, chymotrypsin, SV8 Protease and subtilisin along with inhibitors to generate small active fragments for crystal screening. (Aled Edwards, McMaster University, CANADA & Brian McKeever, Merck, USA). 

Try the Proti-Ace kits from Hampton Research.

Perform screens in duplicate at both 4 degrees Celsius and room temperature. If you notice a difference in solubility between the two temperatures be sure to include temperature as a variable during optimization.

Not enough sample to perform screens in duplicate at two different temperatures? Set up screens at room temperature. After four weeks move the plates to 4°C and watch for changes in solubility. Notice a difference? Use temperature as a variable during optimization.

Remember when using volatile organics as the precipitating agent that no reservoir needs to be added to the drop since distillation and equilibration proceed from the reservoir to the drop.

No crystals, only precipitate in initial screens? Bring precipitant/protein concentration to just below supersaturation and vary pH and temperature to manipulate sample solubility.

Sample soluble at low ionic strength but forms a precipitate when dialyzed against water? This "salting in" effect can be used as a method to grow crystals. Begin with the sample in low ionic strength and dialyze the sample against a solution of lower ionic strength. Or, try changing the pH or temperature with the sample at a constant ionic strength. Charge distribution, surface features, or conformation may change as a function of these variables which in turn may influence sample solubility and the ability to crystallize the sample.

Uncertain about the ligand requirements for your sample? If you have the amino acid sequence for your protein one might consider a sequence homology search to see if the sample is !structurally related! to other proteins with known ligand binding. Then try screening crystallization conditions including the ligands identified through the homology search.

If you are having trouble reproducing crystallization results initially found using small drops (100 nl plus 100 nl for example) and are having trouble growing the crystals in the same conditions in larger drops (1 ul plus 1 ul for example) one might consider the following tips. 

Heather Ringrose (Pfizer UK) has suggested that if you initially screen with 100 nl (reservoir) + 200 nl (protein) you will pick up conditions that scale up to 1 + 1 microlitres. 

Patrick Shaw-Stewart (Douglas Instruments, UK) reported that data-mining of published high-throughput results suggests that increasing the salt can help when scaling up from nanodrops. Presumably this speeds up equilibration. Patrick suggests that if you have a hit where the main precipitant is salt, try increasing the salt concentration in the reservoir solution by 1 M or more. Also, If you have a hit where the main precipitant is PEG or another organic, try increasing reservoir salt concentration by, say, 200 mM. 

If you observe more precipitate in the larger drops compared to the smaller drops, try decreasing the amount of protein by 30 to 60% in the larger drop. In a small drop, the percentage of drop exposed to plastic and air is much greater than a larger drop and protein may adhere to the plastic or be lost at the drop/air interface. To compensate for this potential loss, when moving to a larger drop, reduce the amount of protein used in the large drop to 30 to 60% of that used in the smaller drop. For example, if you used 100 nl of 10 mg/ml protein plus 100 nl reagent, try 1 ul of 5 mg/ml plus 1 ul of reagent.

Should one use an old crystallization screen kit, one past its best of used by date, or make screen and optimization reagents with old stock solutions? 

The main issue would be the ability to reproduce any crystallization results achieved with the old reagents. With time, even at 4 and -20 degrees Celsius the pH, ionic strength, concentration and stability of the reagents will change. Reagents containing polyethylene glycol undergo oxidation which produces increased levels of aldehydes and peroxides which lowers the pH and increases the ionic strength and produces small molecule species which can affect crystallization. Evaporation from the reagents, even when frozen, increases the concentration of solutes and decreases the concentrations of solvents. The result is that one might experience different results with an old reagent compared to a new reagent. An old reagent might not produce crystals as effectively as a new reagent. Or, the old reagent might produce crystals not able to be grown with the new reagent. In such cases it can be difficult or impossible to reproduce or optimize a crystal produced with an old reagent using freshly prepared reagents. However, in these situations, some have reported success using the old reagent as an additive into freshly prepared reagents. For example, mix 1 part old reagent with 9 parts new reagent into the reagent well, mix and then add this reagent to the sample drop. This procedure will incorporate some level of pH, ionic strength and small molecule change from the old reagent into the new reagent. 

The crystallization problems associated with old reagents versus new reagents seems sample dependent. For some samples a change in pH, ionic strength or the presence of a small molecule contaminant produced with age may have no affect on the crystallization. For others the change might be advantageous and for other detrimental. 

The following is a crystallization screening and optimization technique, which can be used in an attempt to improve the quality and size of a crystal or produce a different crystal form. First, one must begin with an initial crystallization reagent (ICR) which has produced some form of a crystal, microcrystalline precipitate or similar promising result. The ICR will be used to bias a subsequent crystallization screening experiment by adding the ICR to the crystallization drop along with the sample and the screen reagent. Pipette the crystallization screen into the crystallization plate reservoir. One may choose to either repeat the original screen or choose a different set of screen reagents. When creating the drop, mix the protein, screen reagent and ICR in a 3:2:1 ratio. 3 parts sample : 2 parts screen reagent : 1 part ICR. The inclusion of the ICR in the drop modifies the crystallization screen, biasing the screen reagents with a promising reagent mix which may increase the likelihood for crystallization, improved crystals, or a different crystal form. 

Possible modifications to this technique include 1) varying the ratio of sample : screen reagent : ICR, and 2) modifying the bias such that one uses the ICR as the screen reagent and instead uses the original or new screen as the bias (3 parts protein : 2 parts ICR : 1 parts screen reagent. Essentially using the crystallization screen as an additive screen. 

References 

The role of bias in crystallization conditions in automated microseeding. F. J. St John, B. Feng and E. Pozharski. Acta Cryst. (2008). D64, 1222-1227. 

Enhancing protein crystallization through precipitant synergy. Majeed S, Ofek G, Belachew A, Huang CC, Zhou T, Kwong PD. Structure. 2003 Sep;11(9):1061-70.

If you observe mostly precipitate in your drops after setting a crystallization screen, dilute the sample 1:1 with sample buffer and repeat the screen. Or, use the original sample concentration and change the sample:reagent drop ratio from 1:1 to 1:2 for the precipitated drops or 2:1 for the clear drops. 

Edwin Pozharski 
University of Maryland, Baltimore

Try adding 2x protein solution + 1x seed solution + 1x reservoir. The reason for doing this is twofold. 


1) The protein/reservoir ration is similar to the condition that generated the seed stock. 

2) A deliberate bias toward the initial condition is provided. 

One might also try 100 nanoliter protein + multi-aspirate mode x (15nl seeds + 85nl reservoir solution). 

One should consider exploring other drop ratios during MMS.