Tables for
Volume F
Crystallography of biological macromolecules
Edited by E. Arnold, D. M. Himmel and M. G. Rossmann

International Tables for Crystallography (2012). Vol. F, ch. 4.1, pp. 107-111   | 1 | 2 |

Section 4.1.4. Advanced crystallization methodologies

C. Sauter,a B. Lorber,b A. McPhersonc and R. Giegéd*

aInstitut de Biologie Moléculaire et Cellulaire (IBMC), Centre National pour la Recherche Scientifique (CNRS), 15 rue René Descartes, Strasbourg, F-67084, France,bUPR 9002, IBMC–CNRS, 15 rue René Descartes Cedex, Strasbourg, 67084, France,cDepartment of Molecular Biology and Biochemistry, University of California, 560 Steinhaus, Irvine, CA 92697–3900, USA, and dMachineries Traductionnelles, ARN, UPR 9002, IBMC du CNRS, 15 rue René Descartes, Strasbourg, 67084, France
Correspondence e-mail:

4.1.4. Advanced crystallization methodologies

| top | pdf |

In methods that manipulate physical parameters, the effects on crystallization are manifold. Among others, they may influence fluid properties in the crystallization media and movement of molecules (gravity), alter the conformation of the macromolecule (pressure), orient crystals (magnetic field), or influence nucleation (electric field). Thus, initiation of crystallization may be triggered by various mechanisms, growth may be differently influenced and, in favourable cases, crystal quality improved. Crystallization in convection-free media

| top | pdf |

Theoretical considerations. When a crystal starts to grow, it attracts surrounding molecules and creates a concentration gradient. Since crystallization occurs on earth in the gravity field, this gradient of concentration and density will lead to convective currents in the mother liquor. In addition, as soon as the crystal becomes big enough, it will sink to the bottom of the solution. Convection and sedimentation almost always take place in classical experimental setups and they almost certainly influence crystallization processes. Their contribution would be drastically reduced in the absence of gravity, as occurs in weightlessness, and the theory predicts more regular crystal growth under a microgravity–diffusive regime that should favour enhanced crystal quality. Such considerations have justified space-crystallization programmes and, as a consequence, have contributed to a deeper understanding of the crystallization process of biomacromolecules (Giegé et al., 1995[link]; McPherson, 1996[link]; Kundrot et al., 2001[link]). However, because of limited access to space experimentation, crystallization in weightlessness will never be user friendly. This has stimulated studies for finding easy ways to simulate microgravity conditions in the laboratory. Such methods, where crystal growth is less dependent on convection and more on diffusion, take advantage of gelled media and microfluidic environments.

Use of microgravity. The first observation in microgravity was that the absence of sedimentation permits the growth of individual crystals in suspension, without any perturbation by contact with vessel walls and neighbouring crystals. However, one should bear in mind that even in microgravity small accelerations can occur owing to vehicle movement and crystal displacement has been recorded (e.g. Lorber et al., 2000[link]). Microgravity experiments require specific instrumentation with dedicated reactors based on current batch, dialysis and vapour-diffusion methods, or on more microgravity-relevant approaches, such as free-interface or counter-diffusion, with crystallization vessels often of rather large size (DeLucas et al., 1994[link]; Giegé et al., 1995[link]; McPherson, 1996[link]; Gonzalez-Ramirez, Carrera et al., 2008[link]). Reproducible data, with a substantial number of model proteins (lysozyme, thaumatin, canavalin and several plant viruses), were obtained in the versatile Advanced Protein Crystallization Facility instrument from the European Space Agency (Vergara et al., 2003[link], 2005[link]). Altogther, an overall positive effect of microgravity on protein crystal growth emerged.

In support of this conclusion are observations of larger sizes for space-grown crystals and improved optical quality, as exemplified by thaumatin crystals (Ng, Lorber et al., 1997[link]). The maximum resolution of diffraction patterns also indicated superiority for microgravity crystals. A striking example is parvalbumin that diffracts to 0.9 Å resolution, while the earth-grown crystals are not suitable for diffraction analysis (Declercq et al., 1999[link]). Also, in several instances, the signal-to-noise ratio of X-ray diffraction data collected from space crystals was greater than for the corresponding earth controls, as for satellite tobacco mosaic virus (McPherson, 1996[link]) and thaumatin (Ng, Lorber et al., 1997[link]; Lorber, Sauter, Robert et al., 1999[link]). An additional criterion is the reduced mosaic spread of reflections recorded from space samples (Snell et al., 1995[link]; Stojanoff et al., 1996[link]; Ng, Lorber et al., 1997[link]; Lorber, Sauter, Robert et al., 1999[link]). Impurity incorporation during growth is another issue and, as shown with lysozyme, the microgravity-grown crystals incorporate fewer impurities than the earth controls (Carter et al., 1999[link]). The best criterion for enhanced crystal quality, however, is the crystallographic structure. In a case study with lysozyme, significant improvement of resolution from 1.6 to 1.35 Å, decreased atomic displacement parameters (ADPs or B factors) and a structure of increased clarity have been noted for the space-grown crystals (Carter et al., 1999[link]). In another study on an aspartyl-tRNA synthetase, a strictly comparative analysis showed that crystals grown in microgravity were superior in many respects to controls prepared under otherwise identical conditions on earth, facilitating structure determination at 2.0 Å resolution (Ng et al., 2002[link]).

Nevertheless divergent conclusions were reached concerning the quality of the X-ray structure, which was shown to be improved (Carter et al., 1999[link]) or unaffected (Vaney et al., 1996[link]) by microgravity. This contradiction may originate from different levels of impurities present in the protein samples and/or from non-identical growth conditions in different hardware. Conceivably, alteration by gravity of fluid properties could affect nucleation. Transport is of importance, because the large sizes of proteins imply that they have low diffusivities. Elimination of fluid convection may, however, dramatically affect the movement and distribution of proteins in the fluid and their transport and absorption to crystal surfaces. In addition, many proteins form non-specific aggregates in solution. These may be a major source of the contaminants that are incorporated into crystal lattices. By virtue of their size and low diffusivity, the movement of aggregates and large impurities in solution is even more significantly altered.

On earth there is continuous density-driven convective mixing in the solution due to gradients arising from temperature or from incorporation of molecules by the growing crystal. The effects of diffusive transport in the laboratory are, by comparison with the microgravity case, almost negligible because of the very slow rate of diffusion of large proteins. Because of convective mixing, protein crystals nucleated on earth are continuously exposed to the full concentration of protein nutrient present in the bulk solvent. Convection thus maintains, at the growing crystal interface, excessive supersaturation as growth proceeds. This provides an explanation as to why microgravity may improve the quality of protein crystals. The mechanism for enhanced order and reduction of defects may not be directly due to convective turbulence at growing crystal surfaces, but to reduction of the concentration of nutrient molecules and impurities in the immediate neighbourhood of the growing crystals. As a protein crystal forms in microgravity, a concentration gradient or `depletion zone' is established around the nucleus. Because protein diffusion is slow and that of impurities may be even slower, the depletion zone is quasi-stable. The net effect is that the surfaces of the growing crystal interface with a local solution phase at a lower concentration of protein nutrient and impurities than exists in the bulk solvent. The crystal, as it grows, experiences a reduction in its local degree of supersaturation and essentially creates for itself an environment equivalent to the metastable region where optimal growth is expected.

Investigations of protein crystallization diverged along two paths. The objective of the first was to produce high-quality crystals for X-ray diffraction analysis. The crystals themselves were the product of the space-bound experiment and biochemical results were secondary. The goal of the second line of investigation was to understand and to control the physics of the process. This second interest was supported by extensive ground-based research. The confluence of results yielded persuasive explanations for the observed improvements in size and quality of protein crystals grown in microgravity and a robust theoretical framework for understanding the phenomena involved. They showed also that protein crystals are more sensitive to the very high degrees of supersaturation at which they are usually grown and to the mass-transport mechanisms responsible for bringing nutrient to their growing surfaces. The self-regulating nature of protein crystallization in microgravity, through the establishment of local concentration gradients of reduced supersaturation, explains why the diffusive transport that predominates produces a significant difference in ultimate crystal quality.

Crystallization in gelled media. Because convection occurs in free solutions, crystallization in gels represents what is essentially a convection-free environment (Henisch, 1988[link]). Thus, the quality of crystals may be improved in gels. Whatever the mechanism of crystallization in gels, the procedure will produce changes in the nucleation and crystal-growth processes, as has been verified with many proteins (Robert & Lefaucheux, 1988[link]; Cudney et al., 1994[link]; Vidal et al., 1999[link]; Biertümpfel et al., 2002[link]; Lorber et al., 2009[link]). Two types of gels have been used, namely, agarose and silica gels. The latter seem to have proven the most adaptable, versatile and useful for proteins. With both agarose and silica gels, it is possible to use a variety of different crystallants, including salts, organic solvents and polymers such as PEG (Gonzalez-Ramirez, Caballero & Garcia-Ruiz, 2008[link]). They also allow the investigator to control pH and temperature. The most successful efforts have involved direct diffusion arrangements, where the crystallant is diffused into a protein-containing gel or vice versa. In practice, experiments are conducted in semi-liquid gelled media where the agarose concentration does not exceed 0.6%(w/v), but crystallization can also take place in 2.0%(w/v) agarose viscoelastic gels, a condition that does not affect the crystal structures (Sugiyama et al., 2009[link]).

Crystals grown in gels can easily be removed from their soft environment and set up for X-ray analysis. They tend to be robust since, as shown with lysozyme crystals, agarose fibres are incorporated into the crystal lattice (Gavira & Garcia-Ruiz, 2002[link]). Gel growth, because it suppresses convection, has proven to be a useful technique for analysing concentration gradients around growing crystals by interferometric techniques (Robert et al., 1994[link]) and growth mechanisms by differential interference contrast microscopy (Van Driessche, Otalora, Gavira & Sazaki, 2008[link]). In conclusion, gelled media mimic microgravity conditions, preserve crystals once they are grown and, as expected, crystals grown in gels are often of superior quality to controls grown from solutions (Zhu et al., 2001[link]; Moreno et al., 2002[link]; Sauter et al., 2002[link]). Finally, gels can prevent damage during crystal soaking and cryo-cooling (Biertümpfel et al., 2005[link]; Sauter et al., 2009[link]; Lorber et al., 2009[link]).

Crystallization in microfluidic devices. Microfluidic devices were recently introduced in the field of biological crystal growth and represent a new means of crystallizing under diffusive conditions. These systems were primarily intended to miniaturize and to parallelize crystallization assays, thus leading to novel, cost-effective, high-throughput screening approaches. However, because of the small size of their channels and chambers (typically below 100 µm in depth and width), they also provide a diffusive environment comparable to that existing in a capillary tube, in a gel, or under microgravity.

Indeed, the first microfluidic application in biocrystallization was a miniaturized version of the free-interface diffusion technique in which the absence of convection is essential (Hansen et al., 2002[link]). The chip consists of a complex integrated fluidic circuit including two networks of channels, one for liquid handling and a second serving as actuation valves. The chip is dedicated to high-throughput screening and, in its initial version, was designed to test 48 crystallization conditions with less than 10 µl of sample solution. Three parallel sets of chambers are used to bring into contact different proportions of macromolecule and crystallant solutions (Fig.[link]a). This concept of chip was further modified to combine free-interface diffusion with vapour diffusion (or vapour permeation) for fine tuning the supersaturation achieved in crystallization chambers (Hansen et al., 2006[link]). This technology also led to a `formulator chip' that can perform hundreds of mixing operations in just a few hours in order to establish precipitation diagrams (Fig.[link]b). A single assay consumes less than 5 nl sample/buffer/precipitant solution and derived precipitation maps are used to delineate a grid of conditions for crystallization screening (Hansen et al., 2004[link]).


Figure | top | pdf |

Examples of microfluidic devices designed for biocrystallization. (a) Schematic view of one of the 46 modules composing the free-interface diffusion chip. An integrated fluidic circuit dispenses the macromolecule and crystallant solutions into the final chambers. The microchannels are then closed and those connecting the top and bottom chambers are opened with the help of pneumatic valves integrated in the chip. The crystallant diffuses in the protein chamber and triggers crystallization. In this version of the chip, each module is divided into three pairs of chambers with volumes of 5–20 nl to create different protein-to-crystallant concentration ratios. (b) The mixing rotor of a formulation chip designed for the high-throughput study of precipitation diagrams. This chip generates protein/buffer/crystallant mixtures at different concentrations in its 5 nl rotor. The three valves in a row constitute a peristaltic pump that homogenizes the mixture. A charge-coupled device (CCD) camera is used to detect the appearance of a precipitate. (c) The nanobatch chip. Nanodroplets are produced in a microfluidic channel and displaced by inert oil (the flow rate determines the drop size from 10 to 20 nl). Droplets can be stored in the chip or in capillaries connected to the exit of the chip. They can be inspected and crystals can be characterized by X-ray diffraction. (d) Microfluidic chip for counter-diffusion experiments. This method relies on the diffusion of a crystallant into an elongated chamber (the microfluidic channel) containing a macromolecular solution. A concentration gradient is generated that develops along the entire crystallization chamber. The propagating supersaturation wave of gradually decreasing amplitude tests a broad range of nucleation and growth conditions in a single experiment. While a precipitate may form at the entrance of the chamber, monocrystals may grow at the opposite end. Crystals can be observed and analysed by X-ray diffraction directly inside the chip (adapted from Sauter et al., 2007[link]).

The second crystallization method implemented in microfluidics was `batch in nanodroplets' (Zheng et al., 2003[link]). The chip design is extremely simple: it consists of inlets for protein, buffer and crystallant solutions, and a microfluidic channel in which 10 nl droplets are prepared by mixing these solutions in various ratios. This device allows a daily formulation of thousands of nanodrops or plugs (Fig.[link]c), which are carried by a flow of inert fluorocarbon oil. They are stored on the chip or in capillary tubes plugged at the exit of the chip and their content can easily be analysed by X-ray diffraction (Yadav et al., 2005[link]). This method is very well suited for high-throughput screening and, in addition to crystal growth, this technology can be used for many applications in chemistry (Song et al., 2006[link]).

Based on the nanodrop approach, a more complex system has recently been designed for basic research purposes. It is able to formulate droplets and to flow them to storage chambers where they can be concentrated or diluted by water permeation through the chambers' walls. This `phase chip' is designed to establish phase diagrams with total control over supersaturation, nucleation and growth kinetics in each individual drop (Shim et al., 2007[link]).

As for free-interface diffusion, the absence of convection in microfluidic channels makes microsystems very appealing for implementing counter-diffusion experiments (Sauter et al., 2007[link]; Ng et al., 2008[link]). Characteristic counter-diffusion features were successfully reproduced in microchannels with a production of crystalline material ranging from microcrystals to large monocrystals along the supersaturation gradient. When made of appropriate polymer material, these counter-diffusion chips also allow a direct on-chip characterization of the crystals by X-ray diffraction, without any further (and potentially deleterious) sample handling (Ng et al., 2008[link]; Dhouib et al., 2009[link]).

These examples illustrate the many advantages of microfluidic chips – low sample consumption, high-throughput screening capabilities, quasi-ideal convectionless growth conditions – and one can anticipate that microfluidic technology will become a popular and affordable tool both for condition screening, optimization and X-ray analysis, and for basic crystallogenesis research.

Simulating other aspects of microgravity crystal growth. Heterogeneous nucleation or crystal growth on the solid surface of crystallization vessels can be avoided under levitation (Rhim & Chung, 1990[link]) and more easily in batch between two oil layers (Chayen, 1996[link]; Lorber & Giegé, 1996[link]). This can also be achieved for the growth of large protein crystals by mild stirring of the solution in two-liquid systems (Adachi et al., 2004[link]).

It was conjectured that other features of weightlessness, such as suppression of convection, could be achieved in the laboratory under hypergravity and when magnetic or electric fields are applied. These possibilities have been tested experimentally. Crystals were grown under forced diffusive transport of the macromolecules in centrifuges (Karpukhina et al., 1975[link]; Lenhoff et al., 1997[link]; Lorber, 2008[link]) and nucleation was shown to be affected by magnetic (Ataka et al., 1997[link]; Sazaki et al., 1997[link]) or external electric (Taleb et al., 1999[link]; Nanev & Penkova, 2001[link]) fields. Interestingly, under these last conditions, growing crystals were shown to have preferential orientations and specific spatial distributions in the crystallization chambers. Magnetic fields produced by small permanent magnets of 1.25 T are sufficient to produce these effects (Astier et al., 1998[link]) and numerical predictions revealed that magnetization forces could damp convection (Qi et al., 2001[link]). For crystallization induced by electric fields, simple devices adapted to vapour-diffusion (Charron et al., 2003[link]) and batch (Al-Haq et al., 2007[link]) methods are available. Crystallization can also be electrochemically assisted by internal electric fields (Frontana-Uribe & Moreno, 2008[link]). In some cases, magnetic and electric fields have been coupled and experiments conducted in gelled media (Sazaki et al., 2004[link]; Moreno et al., 2009[link]).

Although the above methods are not widespread and the underlying physics not completely validated, they can be useful in special cases. For instance, when crystallization attempts systematically yield showers of microcrystals, crystallization inside electric or magnetic fields can be an alternative to obtain monocrystals suitable for X-ray data collection, because the number of nucleation sites is reduced and can be controlled (Moreno & Sazaki, 2004[link]; Hammadi et al., 2009[link]) and crystal quality maintained (Sato et al., 2000[link]; Lübbert et al., 2004[link]). Methods making use of temperature and pressure

| top | pdf |

Temperature and pressure are familiar thermodynamic parameters. Indeed, many living organisms are thermophiles, even hyperthermophiles, or barophiles/piezophiles and thus have evolved macromolecules stable at temperatures up to 110 °C or pressures up to 100 MPa, i.e. 1000-fold atmospheric pressure (Abe & Horikoshi, 2001[link]). Temperature can trigger nucleation, regardless of the crystallization method. This can be done in a controlled manner, but often occurs as an unexpected consequence of accidental temperature variation in the laboratory. Dedicated systems have been designed for temperature-dependent control of nucleation and growth (Astier & Veesler, 2008[link]), and find application for, among other things, the growth of large high-quality protein crystals for neutron crystallography (Budayova-Spano et al., 2007[link]).

Pressure, as anticipated, can trigger nucleation and sustain protein crystal growth (e.g. Suzuki et al., 2002[link]). To facilitate analysis of crystallization output, assays under pressure can be done in agarose gel (Kadri et al., 2003[link]). Rather simple equipment is required allowing batch crystallization of [\sim]12 individual samples of [\sim]80 µl that can be collectively pressurized up to [\sim]400 MPa (Lorber et al., 1996[link]). The effects exerted by pressure are multiple and protein dependent, with habit, number, length, shape and solubility of crystals modified under pressure. Further, crystallization volumes and diffraction properties are affected and, interestingly, these physical properties are essentially conserved upon depressurization of the crystals. In particular, differences in the water sites surrounding thaumatin crystals grown at 0.1 and 150 MPa have been observed (Charron et al., 2002[link]). Crystallographic analysis of cowpea mosaic virus crystals compressed at 330 MPa in a diamond-anvil cell demonstrated pressure-induced ordering of the crystals, lower ADPs and a larger number of ordered water molecules (Girard et al., 2005[link]; Lin et al., 2005[link]). Methods making use of crystallization chaperones

| top | pdf |

Another strategy that has been used for recalcitrant proteins is to combine them in some manner with a second protein, sometimes called a cocrystallization or chaperone protein (Warke & Momany, 2007[link]; Koide, 2009[link]), so that the complex of the two provides an additional chance for success. The idea was first tested with lysozyme complexed with an Fab antibody fragment (Boulot et al., 1988[link]) and has been used particularly with membrane proteins and antibody domains directed against the target protein (Ostermeier et al., 1995[link]). In those cases the antibody fragment enhanced the solubility of the otherwise hydrophobic protein and provided additional lattice contacts in the resultant crystals. There is, in principle, no reason why such `crystallization chaperones' could not be used with soluble proteins. Likewise the method can be useful for the crystallization of functional RNA fragments (Ye et al., 2008[link]). An alternate possibility with great potential is the recently developed DARPin technology based on the natural ankyrin repeat protein fold with randomized surface residue positions allowing specific binding to virtually any target protein (Sennhauser & Grütter, 2008[link]). Seeding

| top | pdf |

It is often necessary to reproduce crystals grown previously, where either the formation of nuclei is limiting, or spontaneous nucleation occurs at such a profound level of supersaturation that poor growth results. In such cases, it is desirable to induce growth in a directed fashion at low levels of supersaturation. This can be accomplished by seeding a metastable, supersaturated protein solution with crystals from earlier trials. Seeding also permits one to uncouple nucleation and growth. Seeding techniques fall into two categories that employ either macroseeds (Thaller et al., 1985[link]) or microcrystals as seeds (Stura et al., 1999[link]; Bergfors, 2003[link]). In both cases, the solution to be seeded should be only slightly supersaturated so that controlled growth can occur.

When seeding with crystals large enough to be manipulated under a microscope, the most important consideration is to eliminate spurious nucleation by transfer of too many seeds. This drawback may be overcome using laser tweezers, a technique that permits non-mechanical, in situ manipulation of individual seeds as small as 1 µm (Bancel et al., 1998[link]). Even if a single large crystal is employed, microcrystals adhering to its surface may be carried across to the fresh solution. To avoid this, the macroseed is washed by passing it through a series of intermediate transfer solutions. In doing so, not only are microcrystals removed, but if the wash solutions are chosen properly, some limited dissolution of the seed surface may take place. This has the effect of freshening the seed-crystal surfaces and promoting new growth once it is introduced into the new protein solution. Note that crystals of homologous macromolecules can serve as seeds (Thaller et al., 1985[link]).

In the second approach with microcrystals, the danger is that too many nuclei will be introduced into the fresh supersaturated solution, and masses of crystals will result. To overcome this, a stock solution of microcrystals is serially diluted over a very broad range. Some dilution sample in the series will, on average, have no more than one microseed per ml; others will have several times more, or none at all. An aliquot ([\sim]1 µl) of each sample in the series is then added to fresh crystallization trials. This empirical test, ideally, identifies the correct sample to use for seeding by yielding only one or a small number of single crystals when crystal growth is completed. Microseeds can be introduced into crystallization trials at any stage of microbatch or vapour-diffusion experiments (D'Arcy, MacSweeney & Haber, 2003[link]; D'Arcy et al., 2004[link]) and this process can be automated (D'Arcy et al., 2007[link]; Newman et al., 2008[link]; Khurshid et al., 2010[link]).


Abe, F. & Horikoshi, K. (2001). The biotechnological potential of piezophiles. Trends Biotechnol. 19, 102–108.
Adachi, H., Takano, K., Matsumura, H., Inoue, T., Mori, Y. & Sasaki, T. (2004). Protein crystal growth with a two-liquid system and stirring solution. J. Synchrotron Rad. 11, 121–124.
Al-Haq, M. I., Lebrasseur, E., Choi, W. K., Tsuchiya, H., Torii, T., Yamazaki, H. & Shinohara, E. (2007). An apparatus for electric-field-induced protein crystallization. J. Appl. Cryst. 40, 199–201.
Astier, J.-P. & Veesler, S. (2008). Using temperature to crystallize proteins: a mini-review. Cryst. Growth Des. 8, 4200–4207.
Astier, J.-P., Veesler, S. & Boistelle, R. (1998). Protein crystals orientation in a magnetic field. Acta Cryst. D54, 703–706.
Ataka, M., Katoh, E. & Wakayama, N. I. (1997). Magnetic orientation as a tool to study the initial stage of crystallization of lysozyme. J. Cryst. Growth, 173, 592–596.
Bancel, P. A., Cajipe, V. B., Rodier, F. & Witz, J. (1998). Laser seeding for biomolecular crystallization. J. Cryst. Growth, 191, 537–544.
Bergfors, T. (2003). Seeds to crystals. J. Struct. Biol. 142, 66–76.
Biertümpfel, C., Basquin, J., Birkenbihl, R. P., Suck, D. & Sauter, C. (2005). Characterization of crystals of the Hjc resolvase from Archaeoglobus fulgidus grown in gel by counter-diffusion. Acta Cryst. F61, 684–687.
Biertümpfel, C., Basquin, J., Suck, D. & Sauter, C. (2002). Crystallization of biological macromolecules using agarose gel. Acta Cryst. D58, 1657–1659.
Boulot, G., Guillon, V., Mariuzza, R. A., Poljak, R. J., Riottot, M.-M., Souchon, H., Spinelli, S. & Tello, D. (1988). Crystallization of antibody fragments and their complexes with antigen. J. Cryst. Growth, 90, 213–221.
Budayova-Spano, M., Dauvergne, F., Audiffren, M., Bactivelane, T. & Cusack, S. (2007). A methodology and an instrument for the temperature-controlled optimization of crystal growth. Acta Cryst. D63, 339–347.
Carter, D. C., Lim, K., Ho, J. X., Wright, B. S., Twigg, P. D., Miller, T. Y., Chapman, J., Keeling, K., Ruble, J., Vekilov, P. G., Thomas, B. R., Rosenberger, F. & Chernov, A. A. (1999). Lower dimer impurity incorporation may result in higher perfection of HEWL crystal grown in µg – a case study. J. Cryst. Growth, 196, 623–637.
Charron, C., Didierjean, C., Mangeot, J.-P. & Aubry, A. (2003). The `Octopus' plate for protein crystallization under an electric field. J. Appl. Cryst. 36, 1482–1483.
Charron, C., Robert, M.-C., Capelle, B., Kadri, A., Jenner, G., Giegé, R. & Lorber, B. (2002). X-ray diffraction properties of protein crystals prepared in agarose gel under hydrostatic pressure. J. Cryst. Growth, 245, 321–333.
Chayen, N. E. (1996). A novel technique for containerless protein crystallization. Protein Eng. 9, 927–929.
Cudney, B., Patel, S. & McPherson, A. (1994). Crystallization of macromolecules in silica gels. Acta Cryst. D50, 479–483.
D'Arcy, A., MacSweeney, A. & Haber, A. (2003). Using natural seeding material to generate nucleation in protein crystallization experiments. Acta Cryst. D59, 1343–1346.
D'Arcy, A., Sweeney, A. M. & Haber, A. (2004). Practical aspects of using the microbatch method in screening conditions for protein crystallization. Methods, 34, 323–328.
D'Arcy, A., Villard, F. & Marsh, M. (2007). An automated microseed matrix-screening method for protein crystallization. Acta Cryst. D63, 550–554.
Declercq, J.-P., Evrard, C., Carter, D. C., Wright, B. S., Etienne, G. & Parello, J. (1999). A crystal of a typical EF-hand protein grown under microgravity diffracts X-rays beyond 0.9 Å resolution. J. Cryst. Growth, 196, 595–601.
DeLucas, L. J., Long, M. M., Moore, K. M., Rosenblum, W. M., Bray, T. L., Smith, C., Carson, M., Narayana, S. V. L., Harrington, M. D., Carter, D., Clark, A. D., Nanni, R. G., Ding, J., Jacobomolina, A., Kamer, G., Hughes, S. H., Arnold, E., Einspahr, H. M., Clancy, L. L., Rao, G. S. J., Cook, P. F., Harris, B. G., Munson, S. H., Finzel, B. C., McPherson, A., Weber, P. C., Lewandowski, F. A., Nagabhushan, T. L., Trotta, P. P., Reichert, P., Navia, M. A., Wilson, K. P., Thomson, J. A., Richards, R. N., Bowersox, K. D., Meade, C. J., Baker, E. S., Bishop, S. P., Dunbar, B. J., Trinh, E., Prahl, J., Sacco, A. & Bugg, C. E. (1994). Recent results and new hardware developments for protein crystal growth in microgravity. J. Cryst. Growth, 135, 183–195.
Dhouib, K., Khan Malek, C., Pfleging, W., Gauthier-Manuel, B., Duffait, R., Thuillier, G., Ferrigno, R., Jacquamet, L., Ohana, J., Ferrer, J.-L., Théobald-Dietrich, A., Giegé, R., Lorber, B. & Sauter, C. (2009). Microfluidic chips for the crystallization of biomacromolecules by counter-diffusion and on-chip crystal X-ray analysis. Lab Chip, 9, 1412–1421.
Frontana-Uribe, B. A. & Moreno, A. (2008). On electrochemically assisted protein crystallization and related methods. Cryst. Growth Des. 8, 4194–4199.
Gavira, J. A. & Garcia-Ruiz, J. M. (2002). Agarose as crystallisation media for proteins II: trapping of gel fibres into the crystals. Acta Cryst. D58, 1653–1656.
Giegé, R., Drenth, J., Ducruix, A., McPherson, A. & Saenger, W. (1995). Crystallogenesis of biological macromolecules. Biological, microgravity, and other physico-chemical aspects. Prog. Cryst. Growth Charact. 30, 237–281.
Girard, E., Kahn, R., Mezouar, M., Dhaussy, A. C., Lin, T., Johnson, J. E. & Fourme, R. (2005). The first crystal structure of a macromolecular assembly under high pressure: CpMV at 330 MPa. Biophys. J. 88, 3562–3571.
Gonzalez-Ramirez, L. A., Caballero, A. G. & Garcia-Ruiz, J. M. (2008). Investigation of the compatibility of gels with precipitating agents and detergents in protein crystallization experiments. Cryst. Growth Des. 8, 4291–4296.
Gonzalez-Ramirez, L. A., Carrera, J., Gavira, J. A., Melero-Garcia, E. & Garcia-Ruiz, J. M. (2008). Granada Crystallization Facility-2: a versatile platform for crystallization in space. Cryst. Growth Des. 8, 4324–4329.
Hammadi, Z., Astier, J. P., Morin, R. & Veesler, S. (2009). Spatial and temporal control of nucleation by localized DC electric field. Cryst. Growth Des. 9, 3346–3347.
Hansen, C. L., Classen, S., Berger, J. M. & Quake, S. R. (2006). A microfluidic device for kinetic optimization of protein crystallization and in situ structure determination. J. Am. Chem. Soc. 128, 3142–3143.
Hansen, C. L., Skordalakes, E., Berger, J. M. & Quake, S. R. (2002). A robust and scalable microfluidic metering method that allows protein crystal growth by free interface diffusion. Proc. Natl Acad. Sci. USA, 99, 16531–16536.
Hansen, C. L., Sommer, M. O. & Quake, S. R. (2004). Systematic investigation of protein phase behavior with a microfluidic formulator. Proc. Natl Acad. Sci. USA, 101, 14431–14436.
Henisch, H. K. (1988). Crystals in Gels and Liesegang Rings. Cambridge, MA: Cambridge University Press.
Kadri, A., Jenner, G., Damak, M., Lorber, B. & Giegé, R. (2003). Crystallogenesis studies of proteins in agarose gel – combined effect of high hydrostatic pressure and pH. J. Cryst. Growth, 257, 390–402.
Karpukhina, S. Y., Barynin, V. V. & Lobanova, G. M. (1975). Crystallization of catalase in the ultracentrifuge. Sov. Phys. Crystallogr. 20, 417–418.
Khurshid, S., Haire, L. F. & Chayen, N. E. (2010). Automated seeding for the optimization of crystal quality. J. Appl. Cryst. 43, 752–756.
Koide, S. (2009). Engineering of recombinant crystallization chaperones. Curr. Opin. Struct. Biol. 19, 449–457.
Kundrot, C. E., Judge, R. A., Pusey, M. L. & Snell, E. H. (2001). Microgravity and macromolecular crystallography. Cryst. Growth Des. 1, 87–99.
Lenhoff, A. M., Pjura, P. E., Dilmore, J. G. & Godlewski, T. S. Jr (1997). Ultracentrifugal crystallization of proteins: transport-kinetic modelling and experimental behavior of catalase. J. Cryst. Growth, 180, 113–126.
Lin, T., Schildkamp, W., Brister, K., Doerschuk, P. C., Somayazulu, M., Mao, H. K. & Johnson, J. E. (2005). The mechanism of high-pressure-induced ordering in a macromolecular crystal. Acta Cryst. D61, 737–743.
Lorber, B. (2008). Virus and protein crystallization under hypergravity. Cryst. Growth Des. 8, 2964–2969.
Lorber, B. & Giegé, R. (1996). Containerless protein crystal­lization in floating drops: application to crystal growth monitoring under reduced nucleation conditions. J. Cryst. Growth, 168, 204–215.
Lorber, B., Jenner, G. & Giegé, R. (1996). Effect of high hydrostatic pressure on the nucleation and growth of protein crystals. J. Cryst. Growth, 158, 103–117.
Lorber, B., Ng, J. D., Lautenschlager, P. & Giegé, R. (2000). Growth kinetics and motion of thaumatin crystals during USML-2 and LMS microgravity missions and comparison with earth controls. J. Cryst. Growth, 208, 665–677.
Lorber, B., Sauter, C., Robert, M.-C., Capelle, B. & Giegé, R. (1999). Crystallization within agarose gel in microgravity improves the quality of thaumatin crystals. Acta Cryst. D55, 1491–1494.
Lorber, B., Sauter, C., Théobald-Dietrich, A., Moreno, A., Schellenberger, P., Robert, M.-C., Capelle, B., Sanglier, S., Potier, N. & Giegé, R. (2009). Crystal growth of proteins, nucleic acids, and viruses in gels. Prog. Biophys. Mol. Biol. 101, 13–25.
Lübbert, D., Meents, A. & Weckert, E. (2004). Accurate rocking-curve measurements on protein crystals grown in a homogeneous magnetic field of 2.4 T. Acta Cryst. D60, 987–998.
McPherson, A. (1996). Macromolecular crystal growth in microgravity. Crystallogr. Rev. 6, 157–305.
Moreno, A., Saridakis, E. & Chayen, N. E. (2002). Combination of oils and gels for enhancing the growth of protein crystals. J. Appl. Cryst. 35, 140–142.
Moreno, A. & Sazaki, G. (2004). The use of a new ad hoc growth cell with parallel electrodes for the nucleation control of lysozyme. J. Cryst. Growth, 264, 438–444.
Moreno, A., Yokaichiya, F., Dimasi, E. & Stojanoff, V. (2009). Growth and characterization of high-quality protein crystals for X-ray crystallography. Ann. N. Y. Acad. Sci. 1161, 429–436.
Nanev, C. N. & Penkova, A. (2001). Nucleation of lysozyme crystals under external electric and ultrasonic fields. J. Cryst. Growth, 232, 285–293.
Newman, J., Pham, T. M. & Peat, T. S. (2008). Phoenito experiments: combining the strengths of commercial crystallization automation. Acta Cryst. F64, 991–996.
Ng, J. D., Clark, P. J., Stevens, R. C. & Kuhn, P. (2008). In situ X-ray analysis of protein crystals in low-birefringent and X-ray transmissive plastic microchannels. Acta Cryst. D64, 189–197.
Ng, J. D., Lorber, B., Giegé, R., Koszelak, S., Day, J., Greenwood, A. & McPherson, A. (1997). Comparative analysis of thaumatin crystals grown on earth and in microgravity. Acta Cryst. D53, 724–733.
Ng, J. D., Sauter, C., Lorber, B., Kirkland, N., Arnez, J. & Giegé, R. (2002). Comparative analysis of space-grown and earth-grown crystals of an aminoacyl-tRNA synthetase: space-grown crystals are more useful for structural determination. Acta Cryst. D58, 645–652.
Ostermeier, C., Iwata, S., Ludwig, B. & Michel, H. (1995). F-V fragment mediated crystallization of the membrane-protein bacterial cytochrome-C-oxidase. Nat. Struct. Biol. 2, 842–846.
Qi, J., Wakayama, N. I. & Ataka, M. (2001). Magnetic suppression of convection in protein crystal growth processes. J. Cryst. Growth, 232, 132–137.
Rhim, W.-K. & Chung, S. K. (1990). Isolation of crystallizing droplets by electrostatic levitation. Methods, 1, 118–127.
Robert, M.-C., Bernard, Y. & Lefaucheux, F. (1994). Study of nucleation-related phenomena in lysozyme solutions. Application to gel growth. Acta Cryst. D50, 496–503.
Robert, M.-C. & Lefaucheux, F. (1988). Crystal growth in gels: principle and applications. J. Cryst. Growth, 90, 358–367.
Sato, T., Yamada, Y., Saijo, S., Hori, T., Hirose, S., Tanaka, N., Sazaki, G., Nakajima, K., Igarashi, N., Tanaka, M. & Matsuura, Y. (2000). Enhancement in the perfection of orthorhombic lysozyme crystals grown in a high magnetic field (10 T). Acta Cryst. D56, 1079–1083.
Sauter, C., Balg, C., Moreno, A., Dhouib, K., Théobald-Dietrich, A., Chênevert, R., Giegé, R. & Lorber, B. (2009). Agarose gel facilitates enzyme crystal soaking with a ligand analog. J. Appl. Cryst. 42, 279–283.
Sauter, C., Dhouib, K. & Lorber, B. (2007). From macrofluidics to microfluidics for the crystallization of biological macromolecules. Cryst. Growth Des. 7, 2247–2250.
Sauter, C., Lorber, B. & Giegé, R. (2002). Towards atomic resolution with crystals grown in gel: the case of thaumatin seen at room temperature. Proteins, 48, 146–150.
Sazaki, G., Moreno, A. & Nakajima, K. (2004). Novel coupling effects of the magnetic and electric fields on protein crystallization. J. Cryst. Growth, 262, 499–502.
Sazaki, G., Yoshida, E., Komatsu, H., Nakada, T., Miyashita, S. & Watanabe, K. (1997). Effects of a magnetic field on the nucleation and growth of protein crystals. J. Cryst. Growth, 173, 231–234.
Sennhauser, G. & Grütter, M. G. (2008). Chaperone-assisted crystallography with DARPins. Structure, 16, 1443–1453.
Shim, J. U., Cristobal, G., Link, D. R., Thorsen, T., Jia, Y., Piattelli, K. & Fraden, S. (2007). Control and measurement of the phase behavior of aqueous solutions using microfluidics. J. Am. Chem. Soc. 129, 8825–8835.
Snell, E. H., Weisgerber, S., Helliwell, J. R., Weckert, E., Hölzer, K. & Schroer, K. (1995). Improvements in lysozyme protein crystal perfection through microgravity growth. Acta Cryst. D51, 1099–1102.
Song, H., Chen, D. L. & Ismagilov, R. F. (2006). Reactions in droplets in microfluidic channels. Angew. Chem. Int. Ed. Engl. 45, 7336–7356.
Stojanoff, V., Snell, E. F., Siddons, D. P. & Helliwell, J. R. (1996). An old technique with a new application: X-ray topography of protein crystals. Synchrotron Radiat. News, 9, 25–26.
Stura, E. A., Charbonnier, J.-B. & Taussig, M. J. (1999). Epitaxial jumps. J. Cryst. Growth, 196, 250–260.
Sugiyama, S., Tanabe, K., Hirose, M., Kitatani, T., Hasenaka, H., Takahashi, Y., Adahi, H., Takano, K., Murakami, S., Mori, Y., Inoue, T. & Matsumura, S. (2009). Protein crystallization in agarose gel with high strength: developing an automated system for protein crystallographic processes. Jpn. J. Appl. Phys. 48, 075502.
Suzuki, Y., Sazaki, G., Miyashita, S., Sawada, T., Tamura, K. & Komatsu, H. (2002). Protein crystallization under high pressure. Biochim. Biophys. Acta, 1595, 345–356.
Taleb, M., Didierjean, C., Jelsch, C., Mangeot, J.-P., Capelle, B. & Aubry, A. (1999). Crystallization of proteins under an external electric field. J. Cryst. Growth, 200, 575–582.
Thaller, C., Eichele, G., Weaver, L. H., Wilson, E., Karlsson, R. & Jansonius, J. N. (1985). Seed enlargement and repeated seeding. Methods Enzymol. 114, 132–135.
Van Driessche, A. E. S., Otalora, F., Gavira, J. A. & Sazaki, G. (2008). Is agarose an impurity or an impurity filter? In situ observation of the joint gel/impurity effect on protein crystal growth kinetics. Cryst. Growth Des. 8, 3623–3629.
Vaney, M. C., Maignan, M., Riès-Kautt, M. & Ducruix, A. (1996). High-resolution structure (1.33 Å) of a HEW lysozyme tetragonal crystal grown in the APCF apparatus. Data and structural comparison with a crystal grown under microgravity from Spacehab-01 mission. Acta Cryst. D52, 505–517.
Vergara, A., Lorber, B., Sauter, C., Giegé, R. & Zagari, A. (2005). Lessons from crystals grown in the Advanced Protein Crystallization Facility for conventional crystallization applied to structural biology. Biophys. Chem. 118, 102–112.
Vergara, A., Lorber, B., Zagari, A. & Giegé, R. (2003). Physical aspects of protein crystal growth investigated with the Advanced Protein Crystallization Facility in reduced-gravity environments. Acta Cryst. D59, 2–15.
Vidal, O., Robert, M.-C., Arnoux, B. & Capelle, B. (1999). Crystalline quality of lysozyme crystals grown in agarose and silica gels studied by X-ray diffraction techniques. J. Cryst. Growth, 196, 559–571.
Warke, A. & Momany, C. (2007). Addressing the protein crystallization bottleneck by cocrystallization. Cryst. Growth Des. 7, 2219–2225.
Yadav, M. K., Gerdts, C. J., Sanishvili, R., Smith, W. W., Roach, L. S., Ismagilov, R. F., Kuhn, P. & Stevens, R. C. (2005). In situ data collection and structure refinement from microcapillary protein crystallization. J. Appl. Cryst. 38, 900–905.
Ye, J. D., Tereshko, V., Frederiksen, J. K., Koide, A., Fellouse, F. A., Sidhu, S. S., Koide, S., Kossiakoff, A. A. & Piccirilli, J. A. (2008). Synthetic antibodies for specific recognition and crystallization of structured RNA. Proc. Natl Acad. Sci. USA, 105, 82–87.
Zheng, B., Roach, L. S. & Ismagilov, R. F. (2003). Screening of protein crystallization conditions on a microfluidic chip using nanoliter-size droplets. J. Am. Chem. Soc. 125, 11170–11171.
Zhu, D.-W., Lorber, B., Sauter, C., Ng, J. D., Bénas, P., Le Grimellec, C. & Giegé, R. (2001). Growth kinetics, diffraction properties and effect of agarose on the stability of a novel crystal form of Thermus thermophilus aspartyl-tRNA synthetase-1. Acta Cryst. D57, 552–558.

to end of page
to top of page