Comparative Effects of Salt, Organic, and Polymer Precipitants on

Comparative Effects of Salt, Organic, and Polymer Precipitants on Protein Phase Behavior and Implications for Vapor Diffusion Andre´ C. Dumetz,† Aaron M. Chockla,‡ Eric W. Kaler,*,§ and Abraham M. Lenhoff* Center for Molecular and Engineering Thermodynamics, Department of Chemical Engineering, UniVersity of Delaware, Newark, Delaware 19716

CRYSTAL GROWTH & DESIGN 2009 VOL. 9, NO. 2 682–691

ReceiVed October 1, 2007; ReVised Manuscript ReceiVed December 6, 2008

ABSTRACT: Salts, polymers, and organic precipitants commonly used to crystallize proteins share the ability to induce attractive protein-protein interactions, and eventually lead to the formation of crystals. Their effects on protein phase behavior are investigated here. First, the ovalbumin phase diagram at pH 7 was determined separately in ammonium sulfate, poly(ethylene glycol) (PEG) 8000, and 2-methyl-2,4-pentanediol (MPD) solutions. Increasing concentrations of each of these three additives lead to well-defined phase separations that have the same characteristic trends, but differ in their physical appearance. Second, the phase diagrams of ovalbumin in ammonium sulfate and ribonuclease A in MPD were determined at pH 6, conditions under which the two proteins crystallize. The general shape of the phase diagram and the formation kinetics of the different phases are similar for both proteins, suggesting that the main characteristics of the phase behavior are independent of the physical origin of the attraction between protein molecules. Finally, the phase diagrams are compared to the results of vapor diffusion experiments carried out under similar solution conditions. These results illustrate how vapor diffusion experiments can be interpreted in terms of the phase diagram, and how crystallization conditions can be designed based on knowledge of the experimental phase diagrams of proteins. Introduction The precipitants used to crystallize proteins are usually classified as salts, polymers, or organic additives, but these different classes of precipitants have the common property of affecting protein interactions, and under certain conditions lead to crystallization. The second osmotic virial coefficient (b2), which is a direct measure of protein interactions, has been used to investigate the effects of salts,1-8 polymers,9-12 and organic additives,13-17 and it has been demonstrated that changes in phase behavior correspond to increasingly attractive interactions. The goal here is to compare the phase behavior obtained for these three classes of precipitant and to show how the information contained in the phase diagram can be used to design vapor diffusion experiments. Salt and organic precipitants have been used to purify proteins since the early investigations of protein solutions.18,19 Crystallization, which was often achieved unintentionally during purification, was then exploited as a purification step. Subsequent interest in protein crystallization has been driven primarily by the necessity of obtaining protein crystals for structural biology applications, but the fundamental approach originally developed to crystallize proteins as well as the different types of precipitants, that is, salts and organic additives, remain largely the same. Although polymers have been introduced more recently as an additional class of precipitants,20-23 the main improvements in protein crystallization have probably been in the development of vapor diffusion techniques that permit progressive equilibration, and of high throughput methods that permit extensive screening of solution conditions.24,25 Despite the similarities in their use and effects, the different classes of precipitant act in different ways. Good salt precipitants * Authors to whom correspondence should be addressed. E-mail: lenhoff@ udel.edu (A.M.L.), [email protected] (E.W.K.). † Current address: GlaxoSmithKline, 709 Swedeland Road, King of Prussia, PA 19406, USA. ‡ Current address: Department of Chemical Engineering, University of Texas, Austin, TX 78712, USA. § Current address: Office of the Provost and Senior Vice-President, 407 Administration Building, Stony Brook University, Stony Brook, NY 11794, USA.

such as ammonium sulfate are known to be excluded from the protein surface,26,27 and because of the direct correlation between their efficiency and their strength of hydration, protein saltingout has typically been attributed to water-mediated effects.19,28-30 A competition for waters of hydration between salt and protein molecules is not, however, the only effect of the salt, as evidence suggests that ion binding sometimes plays an important role by creating an effective repulsion26,31,32 or by mediating attractive interactions between proteins through involvement in crystal contacts.33-36 Polymers are the most recently introduced class of precipitants, and PEG, the mode of action of which is well established, may be the most widely used of all participants.37-39 PEG is generally preferentially excluded from the protein surface,40-42 and in view of the success of theoretical models43-46 in describing the main experimental trends,10,11,47,48 its effect on protein interactions is commonly attributed to depletion forces. However, this picture is also nuanced by experimental evidence showing that PEG sometimes interacts with proteins.12,49-52 Like salts, organic additives have long been used to purify proteins by precipitation or crystallization.18,19,53 Some are particularly efficient, and for example, 2-methyl-2,4-pentanediol (MPD) alone is one of the most successful additives used to crystallize proteins.37-39 It was originally argued that organic additives affect protein interactions by changing the solvent dielectric constant,18,19,54,55 implying that organic additives affect protein interactions through electrostatic interactions in a similar way to decreasing the salt concentration. However, the effect of an organic solvent on the solution dielectric constant is too weak to explain the effects observed.56 Moreover, this mechanism does not explain why organic additives can crystallize proteins that do not salt-in, or proteins far from their isoelectric points. Crystallographic evidence shows that organic molecules such as MPD57 or small alcohols58-60 can bind to the hydrophobic regions of protein surfaces, thereby significantly affecting protein hydration.61,62 However, despite this evidence, organic precipitants such as MPD are generally preferentially excluded from the protein surface,63 and even if this is not the only effect to

10.1021/cg700956b CCC: $40.75  2009 American Chemical Society Published on Web 01/07/2009

Comparative Effects on Protein Phase Behavior

Figure 1. Schematic representation of a protein phase diagram as a function of precipitant and protein concentrations based on Foffi et al.85

which they give rise, their contribution can be attributed to excluded volume effects.64,65 The same type of effect may also explain the liquid-liquid phase separation that occurs in aqueous MPD solutions with increasing salt concentration in the absence of protein.63 However, much remains to be done to develop a molecular understanding of the effects of organic additives on protein interactions. The different phases that are observed experimentally in protein solutions can be classified as crystals, aggregates, dense liquid phases, or gels. However, their appearance varies considerably among additives as well as among proteins, and the present investigation also aims to present a systematic comparison of the appearance of the different phases obtained with different additives. Protein phase behavior has been investigated mainly in salt solutions for lysozyme,66-68 γ-crystallin,69-73 and more recently BPTI.74-76 On the other hand, protein phase behavior in PEG solutions has been studied in detail for urate oxidase,77,78 glucose isomerase,79 and a few other proteins.80-82 Although organic additives are very common for growing crystals for X-ray diffraction, protein phase behavior in the presence of organic additives has not been the subject of extensive investigation. Theoretically, the phase behavior in salt or organic solutions can be explained within a framework used to explain other colloidal behavior (Figure 1), which for short-range attractive interactions predicts a liquid-liquid phase separation that is metastable with respect to crystallization.83-87 This pattern is similar to ones seen experimentally for proteins in aqueous salt solutions, even though the theoretical approaches are based on a simplistic representation of protein interactions that takes into account only their short-range and attractive nature.83-85 Theoretical phase diagrams of the kind presented in Figure 1 are more typically plotted with the abscissa corresponding to the protein concentration and the ordinate representing the reciprocal precipitant concentration, but this convention was reversed here to facilitate the comparisons with experimental data presented below. Phase behavior calculations for colloid/polymer solutions are different in the sense that they are based on mechanistic models seeking to describe depletion forces.87,88 However, proteins often crystallize at polymer concentrations above the polymer overlap concentration (c*) and with polymers that have a radius of gyration larger than the effective size of proteins. Despite a few recent theoretical studies89,90 this situation has not been completely resolved.

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Recently, based on the results of mode coupling theory (MCT), the idea that liquid-liquid phase separation of protein solutions can lead to the formation of a kinetically trapped phase corresponding to a gel phase has received renewed interest84,85 (Figure 1). The predictions of MCT are in good agreement with experimental investigations in salt solutions, and the theory can explain the formation of precipitates, reversible gels, or aggregates.91,92 However, experimentally the identification of the different phases can be confusing. For example, for lysozyme in sodium chloride, the liquid-liquid phase separation previously reported66-68 to occur at high protein and low salt concentrations requires ultracentrifugation to obtain two clear phases separated by a sharp meniscus. At higher sodium chloride and lower lysozyme concentrations, the formation of a precipitate that is not much different in appearance to that corresponding to the liquid-liquid phase separation can be interpreted as a gel or frustrated liquid-liquid phase separation.91 This is due to the very high viscosity of the dense phase in the case of lysozyme liquid-liquid phase separation, which causes extremely slow coalescence of the dense liquid drops. These observations raise questions regarding the physical nature of the phase separation in PEG solutions, which has also been reported as a liquid-liquid phase separation,78,79 and the phase separation that is expected with increasing concentration of organic additives. By investigating in parallel the experimental phase behavior obtained for salt, polymer, and organic additives, the first goal of the present work is to clarify the appearance and the physical nature of phases observed for different additives. First, the ovalbumin phase diagram is determined as a function of the concentration of ammonium sulfate, PEG 8000 or MPD at pH 7, and the phases obtained are compared. Second, the phase diagrams obtained at pH 6 for ovalbumin in ammonium sulfate and ribonuclease A in MPD (conditions under which these two proteins crystallize) are presented to provide a more complete picture of protein phase behavior. Third, these latter results are compared to the results of vapor diffusion experiments to show how the information contained in protein phase diagrams can be used to improve protein crystallization, which is an important goal of this work. Materials and Methods Proteins and Chemicals. Ovalbumin was purified from fresh, singlecomb white Leghorn eggs using previously published procedures.91,93,94 Ribonuclease A (LS003433) was purchased from Worthington Biochemical Corporation (Lakewood, NJ). PEG 8000 (P-4463), MPD (68340), sodium sulfate (A-2939), sodium malonate (M-4795), MES (M-8250), and bis-tris (B-7535) were purchased from Sigma-Aldrich (St. Louis, MO). Sodium phosphate dibasic (S-374), sodium chloride (S-271), sodium acetate (S-374), sodium hydroxide (S-318), and hydrochloric acid (A144-212) were obtained from Fisher Scientific. The buffers used were 5 mM sodium phosphate at pH 7 and 5 mM bis-tris at pH 6. The buffers were prepared using distilled water further purified with a Milli-Q system from Millipore (Billerica, MA), and the pH was adjusted with concentrated solutions of sodium hydroxide and hydrochloric acid. Purification. The proteins used were further purified by ion-exchange ¨ KTA Purifier from GE Healthcare (Piscatchromatography using an A away, NJ). Ion exchange resins SP Sepharose FF and Q Sepharose FF from GE Healthcare, packed in XK13 and XK28 columns, were used for the purification of ribonuclease A and ovalbumin, respectively. After purification, the protein solutions were reconcentrated up to 30-60 mg/mL using an Amicon stirred ultrafiltration cell (model 8200) and a YM10 ultrafiltration membrane from Millipore. The solutions were then dialyzed extensively against deionized water and finally against lowsalt buffer using 10 mL Slide-A-Lyzer cassettes from Pierce Biotechnology (Rockford, IL). The final step was to reconcentrate the protein

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solutions up to 100-150 mg/mL using a 10k Amicon Ultra-4 centrifugal filter device from Millipore. The protein concentration of the final solutions was measured by UV absorbance at 280 nm using a Lambda 4B spectrophotometer from Perkin-Elmer (Norwalk, CT). The extinction 1% coefficients used were E1cm ) 7.35 for ovalbumin and 7.14 for 95 ribonuclease A. Sample Preparation. Each protein sample, of a total volume 200 µL, was prepared in a 0.5 mL tube (05-408-120) from Fisher Scientific by pipetting appropriate amounts of concentrated stock solutions, and the samples were mixed immediately after preparation. The samples for the phase behavior measurements in ammonium sulfate were prepared by pipetting first the high-salt buffer solution, followed by the low-salt buffer and concentrated protein stock solution. The samples for the phase behavior measurements in PEG were prepared by pipetting first the high-salt buffer followed by the lowsalt buffer, the concentrated protein stock solution, and finally the concentrated PEG stock solution. The samples for the phase behavior measurements in MPD were prepared by pipetting first the low-salt buffer followed by the concentrated MPD and protein stock solutions. From each sample, between 7 and 10 µL were pipetted into a 72-well microplate (HR3-087) and covered by paraffin oil (HR3-421) (Hampton Research, Aliso Viejo, CA). The same procedure was reproduced six times so that each column on the microplate had the same composition. These procedures were found to be appropriate to ensure the reproducibility and accuracy of the measurements. The evolution of the phase behavior was recorded every day during the first week and at regular intervals during the subsequent months. A description of the microscopy methods is included elsewhere.91 Vapor diffusion experiments were carried out using VDXm vapor diffusion plates (HR3-108) purchased from Hampton Research and the same stock solutions as before. The top of each reservoir was coated with high vacuum grease to ensure a hermetic seal. In each reservoir 600 µL of precipitant was prepared by pipetting the appropriate amounts of low salt buffer and concentrated precipitant stock solution. Eight microliter drops were prepared on siliconized glass circle cover slides (HR3-239) from Hampton Research by mixing 4 µL of the precipitant in the reservoir with the same volume of concentrated protein stock solution. The cover slides were then set on top of the reservoirs. Different series of experiments were performed at different protein concentrations (5-100 mg/mL) with a precipitant concentration in the reservoir between 1.6 and 2.4 M for ammonium sulfate and between 55% and 77.5% (v/v) for MPD. The vapor diffusion plates were observed every day during the first week and at regular intervals during the subsequent months.

Results and Discussion Comparison of Ovalbumin Phase Behavior in (NH4)2SO4, PEG 8000, and MPD. Figure 2A shows the ovalbumin phase diagram in ammonium sulfate solutions at pH 7. Ovalbumin aggregated around 2-2.2 M ammonium sulfate, and a second aggregation line corresponding to a well-defined transition was observed around 2.4 M ammonium sulfate. The appearance of the aggregates after 20 min in 2.2 M ammonium sulfate is illustrated in Figure 3. The solutions at 43.7 mg/mL and above were cloudy upon preparation of the samples, and this was used to define the second aggregation line. In Figure 3, between the first and second aggregation lines the aggregates corresponded to small gel beads that formed rapidly, and after 20 min the solutions between 25 and 37.4 mg/mL already contained aggregates. At 37.4 mg/mL, the solution was already cloudy, but growing gel beads were still clearly apparent in the solutions at 25 and 31.2 mg/mL, which ultimately also turned completely cloudy. Figure 4 illustrates the phase behavior in 2.2 M ammonium sulfate at slightly lower protein concentrations after a period of 10 days, and in contrast to the situation observed at higher protein concentrations, under those conditions each well contained gel bead-like aggregates, the number and size of which depended on nucleation events. Figure 2B shows ovalbumin phase behavior in PEG 8000 solutions at pH 7 with a background of 0.5 M sodium chloride.

Figure 2. Ovalbumin in (A) ammonium sulfate, 5 mM sodium phosphate, pH 7 (([) first and (]) second aggregation lines), (B) PEG 8000, 0.5 M NaCl, 5 mM sodium phosphate, pH 7 ((b) liquid-liquid phase separation and (---) precipitation line) and (C) MPD, 5 mM sodium phosphate, pH 7 ((9) aggregation line). The dotted lines represent the maximum protein concentration that was investigated at each precipitant concentration.

Ovalbumin phase separated with increasing PEG concentration around 10-20% PEG (m/v). The supernatant corresponded to an upper phase depleted in protein, whereas the extremely viscous lower phase was enriched in protein. In contrast to ovalbumin in ammonium sulfate, which formed gel bead-like aggregates, PEG 8000/ovalbumin phase separation corresponds to a genuine liquid-liquid phase separation. When the system was perturbed mechanically, the dense phase flowed, albeit very slowly; the liquid drops could take half an hour to coalesce under the effect of surface tension. A second transition corresponding to the formation of precipitates in the dense liquid drops was observed at higher PEG concentrations. However, in contrast to the observations



Figure 3. Appearance of ovalbumin solutions with increasing protein concentration, 20 min after preparation in 2.2 M ammonium sulfate, 5 mM sodium phosphate, pH 7. The scale bar represents 0.1 mm.

Figure 5. Ovalbumin phase behavior in 20% PEG 8000, 0.5 M NaCl, 5 mM sodium phosphate, pH 7 as a function of increasing protein concentration. The scale bar represents 0.3 mm.

Figure 4. Appearance of ovalbumin phase behavior with increasing protein concentration, 10 days after preparation, in 2.2 M ammonium sulfate, 5 mM sodium phosphate, pH 7. The scale bar represents 0.3 mm.

for ovalbumin in ammonium sulfate, which corresponds to a well-defined transition in the kinetics of precipitation upon sample preparation, this second transition corresponded to a change in the appearance of the samples, after coalescence of the dense liquid phase, one day after sample preparation. Furthermore, because it was not possible to define the exact location of the transition, it is reported with a dashed line in Figure 2B. Figure 5 illustrates the phase behavior in 20% PEG (m/v) with increasing protein concentration. At this PEG concentration, ovalbumin phase separated even at low protein concentrations, and the formation of a dark precipitate is clearly apparent in the last five wells. The ovalbumin phase diagram in MPD at pH 7 is reported in Figure 2C. With increasing MPD concentration, ovalbumin formed white aggregates that sedimented to the bottom of the 0.5 mL tubes within a day. At high MPD concentration, however, the samples formed what appeared to be a translucent gel phase, but the solutions were not further characterized. Under the microscope, the solutions became progressively more cloudy with increasing protein concentration, but any transition was difficult to identify. The appearance of ovalbumin phase behavior differed considerably among ammonium sulfate, PEG 8000 and MPD, for which gel bead aggregates, liquid-liquid phase separation and white precipitates were observed, respectively. However, despite these differences, the aggregation lines in ammonium sulfate and MPD and the liquid-liquid phase boundary in PEG have the same characteristic shape (Figure 2).

Previous investigations91 suggested that the first aggregation line of ovalbumin in ammonium sulfate, that is, formation of gel bead-like aggregates, corresponds to a frustrated liquid-liquid phase separation that leads to formation of a kinetically trapped phase. The second aggregation line, which corresponds to the formation of a precipitate immediately upon sample preparation, was then interpreted as the spinodal line.91 By analogy, it is possible to identify ovalbumin liquid-liquid phase separation in PEG 8000 and the ovalbumin aggregation line in MPD as the binodal. The results reported here correspond to the phase behavior of the proteins in their native state, and the formation of the different phases was reversible, that is, the proteins in any phase could be redissolved entirely upon addition of low-salt buffer (data not shown). Some additives are known for their tendency to denature proteins, but ammonium sulfate, PEG and MPD are preferentially excluded from the surfaces of proteins and are known to have a stabilizing effect against denaturation.26,27,40-42,63,96 The formation of aggregates upon denaturation corresponds to a fundamentally different problem, although the physical appearance of the aggregates can be very similar to that of those seen here. Despite the diversity in appearance of the different phases observed for these native proteins, there is a clear similarity in the general structure of the phase transitions. This suggests that the general form of the phase behavior is independent of the origin of the physical attraction between proteins and is only due to protein interactions becoming more attractive. This result may be expected from simple theoretical models84,85 (Figure 1), and the comparison of the phase diagrams determined here for ovalbumin in ammonium sulfate, PEG 8000, and MPD appears to verify those principles. Nonetheless, because ovalbumin does not crystallize easily at pH 7 with any of the three different additives used here, the phase behavior reported in this section does not include the solubility line. Ovalbumin crystallized sporadically in ammonium sulfate and PEG 8000 in the experiments reported above, but these observations were insufficient to determine the protein crystal solubilities. The next section describes the phase diagrams of ovalbumin and ribonuclease A at pH 6 in ammonium sulfate and MPD respectively, solution conditions at which these two proteins crystallize easily.93,97,98


Figure 6. Ovalbumin ([) aggregation line and (]) crystal solubility in ammonium sulfate, 5 mM MES, pH 6.

Figure 7. Appearance of ovalbumin phase behavior with increasing protein concentration after 8 days in 2.16 M ammonium sulfate, 5 mM MES, pH 6.

Protein Crystallization in Ammonium Sulfate and MPD Solutions. Figure 6shows ovalbumin crystal solubility and aggregation lines in ammonium sulfate at pH 6. In contrast to its behavior at pH 7, ovalbumin crystallizes easily between pH 4 and 6,91 and the crystal solubility was measured from the solution supernatant. Figure 7 shows the appearance of the phase behavior with increasing protein concentration after 8 days in 2.16 M ammonium sulfate, pH 6, corresponding to a vertical cut of the phase diagram in Figure 6. The wells at 8.2, 16.5, and 27.7 mg/mL were cloudy upon preparation of the samples due to the formation of amorphous aggregates, but within two or three days, needle-like crystals grew from both clear solutions and in the presence of aggregates, and they reached their final size after a few days. In the case of the last three wells, the aggregates that were initially present dissolved gradually, and contributed to the growth of the crystals. The needle-like crystals were smaller at higher protein concentrations, and in the wells at 16.5 and 27.7 mg/mL (Figure 7), ovalbumin crystals have the appearance of spherulites. The physical nature of the precipitate that forms upon sample preparation is not clear; it may correspond to a gel phase similar to that observed at pH 7 under similar conditions. However, it is not possible to distinguish a very viscous liquid phase from a gel phase based solely on visual observations, especially given that the precipitate evidently redissolves, feeding nucleation and growth of the crystals. The phase diagram for ribonuclease A in MPD at pH 6 (Figure 8) is significantly different from that of ovalbumin in

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Figure 8. Ribonuclease A (0) crystal solubility, (9) liquid-liquid phase separation, and (---) immediate precipitation after sample preparation in MPD, 5 mM MES, pH 6.

Figure 9. Detail of the evolution over time of ribonuclease A at a protein concentration of 33.2 mg/mL in 60% MPD (m/v), 5 mM MES, pH 6. The scale bar represents 0.2 mm.

MPD at pH 7 (Figure 2C), despite similarities in their general shapes. With increasing MPD concentration, ribonuclease A underwent a liquid-liquid phase separation around 60% MPD, and at slightly higher MPD concentrations, the solutions turned cloudy upon sample preparation. However, this second transition, represented by a dashed line in Figure 8, was not as well defined as for the second aggregation line of ovalbumin in ammonium sulfate at pH 7 (Figure 2A), because even at lower MPD concentrations the precipitate formed rapidly. It is for this reason that what is referred to as the second aggregation line is represented as a dashed line in Figure 8. In the vicinity of this transition, crystals appeared after only a few hours and reached their maximum size within a couple of days. After a week, crystals were observed in almost the entire region in which the liquid-liquid phase separation was first observed, but ribonuclease A did not crystallize between the solubility line and the liquid-liquid phase boundary. Figure 9 illustrates the coalescence of drops after liquid-liquid phase separation and the kinetics of crystal growth at 33.2 mg/ mL ribonuclease A in 60% MPD. The initial cloudiness disappeared within a few hours, giving rise by coalescence to large liquid drops. In contrast to the white aggregates seen for ovalbumin in MPD (Figure 2C), the physical nature of which was not investigated further, ribonuclease A solutions in MPD (Figure 8) displayed a true liquid-liquid phase separation. After only a few hours, the first crystals appeared in the dense liquid phase close to where the second transition was observed, and they reached their maximum size after a few days. In Figure 9, which corresponds to one of the samples in which crystallization occurred almost immediately, a small crystal is apparent in one



Figure 10. Appearance of ribonuclease A phase behavior in 60% MPD (m/v), 5 mM MES, pH 6, after one month. The scale bar represents 0.3 mm.

Figure 11. Liquid-liquid phase separation observed without protein in MPD solutions with increasing salt concentration of (b) sodium malonate, ([) ammonium sulfate, (2) sodium acetate, and (9) sodium chloride. The experiments were carried out in 5 mM sodium phosphate buffer, pH 7.

of the central drops after 18 h, and it continued to grow. As is apparent from the empty space around the central drop in the micrograph after 30 h, the drop containing the growing crystal increased in size considerably between 18 and 30 h due to coalescence with neighboring drops. The dense liquid phase was less viscous than that previously obtained for ovalbumin in PEG 8000. The drops flowed slowly and recovered their initial shapes in a few minutes upon perturbation. The role of the proteinrich drops in feeding the growth of crystals is also evident from the shrinking size of the remaining liquid drops over time. Figure 10 shows the final appearance of the phase behavior of ribonuclease A in MPD after one month, corresponding to a vertical line at 60% MPD in Figure 8. The first four wells, which correspond to the region below the liquid-liquid phase separation, remained clear even after a prolonged period of time. At 13.3 mg/mL, the liquid-liquid phase separation is still apparent, but above this concentration, it has disappeared in favor of crystals. Close to the liquid-liquid phase separation, a few crystals appeared after a delay of about one week, and their size could reach a few hundred micrometers, whereas in the vicinity of the second transition or precipitation line, a multitude of small crystals nucleated within a couple of hours, and their ultimate size was much smaller. These observations suggest that nucleation occurs only in solutions that first undergo a liquid-liquid phase separation, and the proximity of the second transition, which may be interpreted as the spinodal line, seems to be a determinant for crystal nucleation. These results corroborate the theoretical results of ten Wolde and Frenkel suggesting that liquid-liquid phase separations lower the free-energy barrier to crystal nucleation.99,100 However, ovalbumin in ammonium sulfate at pH 6 crystallizes easily both from clear solution and in the presence of aggregates, and this suggests that, not surprisingly, more than one route is possible for crystal nucleation. The metastability of the liquid-liquid phase separation to crystallizationpreviouslydemonstratedforsalt67,70 andpolymer78,79 solutions is apparent here for ribonuclease A in MPD solutions. This characteristic of protein phase behavior is important as it justifies the suggestion that aggregation or liquid-liquid phase separation corresponds to a well-defined phase transition and that proteins may still crystallize following these events. Aqueous solutions of MPD in the absence of protein also undergo a liquid-liquid phase separation with increasing salt

concentration, and it was suggested previously that colloidal interactions can be affected by the proximity of such a phase separation.101 Figure 11 shows MPD liquid-liquid phase separation obtained in solutions of four salts that are commonly used in crystallization screens, viz. sodium malonate, ammonium sulfate, sodium acetate, and sodium chloride. A low background buffer concentration was added to maintain the pH constant. In the measurements of ovalbumin and ribonuclease A liquid-liquid phase separation in MPD shown earlier (Figures 2 and 8), the buffer was the only added electrolyte, that is, no salt was added. In the case of ovalbumin those conditions correspond to zero salt concentration in Figure 11, and although the ribonuclease A measurements were made with differences in both pH and buffer from those used in preparing Figure 11, it can be assumed that this did not affect the one-phase region at low salt concentrations. The data in Figure 11 show that salt solutions can phase separate with MPD alone at sufficiently high MPD concentrations. Therefore, protein crystallization conditions can be close to the phase boundary, especially when the buffer is a strongly hydrated salt. This observation suggests that particular attention should be given to the buffer concentration when crystallizing proteins using MPD. Comparison between Experimental Phase Diagram and Vapor Diffusion Experiments. The general structure of the protein phase diagram provides useful information that has obvious practical implications for optimization of the solution conditions in crystallization screens.102,103 However, protein crystals grown for X-ray diffraction are typically obtained by vapor diffusion, and this section explores how the phase diagrams obtained at pH 6 for ovalbumin in ammonium sulfate and ribonuclease A in MPD can explain the results of vapor diffusion experiments for these two systems. Vapor diffusion104 is a dynamic procedure in which the initial concentration in the drop can be determined from the stock solution concentrations, and the final precipitant concentration in the drop should be close to the concentration in the reservoir. The latter point assumes that the protein concentration does not significantly affect the vapor equilibrium and that the dilution of the reservoir is negligible. Figure 12 shows the expected evolution of the ammonium sulfate and ovalbumin concentrations in the hanging drops during the vapor diffusion experiments undertaken here. The aggregate and crystal solubilities correspond to the phase diagram previously obtained for ovalbumin in ammonium sulfate


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Figure 12. (Top) Likely evolution of the salt and protein concentrations in a series of hanging drop vapor diffusion experiments for ovalbumin in ammonium sulfate, 5 mM MES, pH 6, compared to ovalbumin (]) crystal solubility and ([) aggregation line. (Bottom) Appearance after one month of ovalbumin phase behavior in this series of hanging drop vapor diffusion experiments. The ammonium sulfate concentrations correspond to those in the reservoirs and should also correspond to the final concentrations in the hanging drops. The scale bar represents 0.5 mm.

Figure 13. (Top) Likely evolution of the salt and protein concentrations in a series of hanging drop vapor diffusion experiments for ribonuclease A in MPD, 5 mM MES, pH 6, compared to ribonuclease A (0) crystal solubility, (9) liquid-liquid phase separation, and (---) precipitation line. (Bottom) Appearance after one month of ribonuclease A phase behavior in this series of hanging drop vapor diffusion experiments in MPD, 5 mM MES, pH 6. The MPD concentrations correspond to those in the reservoirs and should also correspond to the final concentrations in the hanging drops. The scale bar represents 0.5 mm.

at pH 6 (Figure 6). The initial equilibration, which should lead to doubling of the protein and precipitant concentrations in the hanging drops, takes less than a day, and for the last three micrographs at 2.1, 2.2, and 2.4 M ammonium sulfate led to the formation of precipitates. During the subsequent days, crystals grew, with kinetics very similar to those observed in batch crystallization. Figure 12 also illustrates the appearance after one month of the series of hanging drops prepared at different concentrations of ammonium sulfate. The salt concentration that is reported below the micrographs in Figure 12 corresponds to the concentration in the reservoir, and it should also correspond to the final concentration in the hanging drop given the assumptions previously stated. For ammonium sulfate concentrations of 1.6 M, the drops remained clear of aggregates even after one month. Large clusters of needles were obtained at 1.7 and 1.8 M ammonium sulfate, and small spherulites were obtained at 1.9 M ammonium sulfate and above. Figure 13 shows similar results for ribonuclease A in MPD at pH 6. The crystal solubility and the liquid-liquid phase separation correspond to the data in Figure 8, and the tie-lines correspond to the evolution of protein and MPD concentrations in the hanging drops. Figure 13 also illustrates the final appearance after one month of the series of hanging drops at different concentrations of MPD. The initial equilibration on the first day led to a liquid-liquid phase separation in all the drops except the first two. At high MPD concentrations, however, the dense phase had the appearance of a dark precipitate. Over the subsequent days, crystals grew in the wells in which liquid-liquid phase separation was initially observed.

The only exception is the last micrograph in Figure 13, for which the dark precipitate did not change in appearance after formation. Growing crystals appeared first in the hanging drops at 65 and 67.5% MPD, but the largest crystals were obtained near the liquid-liquid phase boundary, for which only a few crystals nucleated and appeared with a delay of about one week. Crystallization did not occur in the drops at lower MPD concentrations even where the phase diagram indicates that the solution is metastable. For both ovalbumin in ammonium sulfate and ribonuclease A in MPD, the results of the vapor diffusion experiments correspond closely to the observations made during batch crystallization experiments (Figures 7, 9, and 10). The results suggest that in order to optimize vapor diffusion experiments the concentration in the reservoir should be as close as possible to the optimum precipitant concentration used for bulk crystallization. The crystals obtained by vapor diffusion were much larger than those by batch crystallization, illustrating why vapor diffusion is widely used to grow crystals for X-ray diffraction. The clusters of needle-like crystals of ovalbumin in ammonium sulfate obtained by vapor diffusion could be three times the size of those obtained under similar solution conditions by batch crystallization. Similarly, the ribonuclease A crystals could be twice as large, but vapor diffusion did not change either the phase behavior or the appearance of the crystals. The optimum ammonium sulfate concentration for growing ovalbumin crystals at pH 6 is around 1.7-1.8 M, and the optimum MPD concentration for growing ribonuclease A


crystals is 60-62.5% (m/v). Both represent restricted ranges of precipitant concentrations, and this explains some of the difficulty of finding conditions conducive to protein crystallization by empirical screening. It also justifies use of the formation of a precipitate as an efficient indicator for optimizing the precipitant concentration. The effect of the initial protein concentration was investigated from 2.5 to 50 mg/mL, but very similar results were obtained independent of the protein concentration. In many crystallization experiments, only small quantities of protein are available, and it is consequently not possible to determine phase diagrams systematically. However, the present results show that there is a practical equivalence between the information obtained by hanging drop vapor diffusion experiments and that contained in the experimental phase diagram. The present results therefore show how to use the protein phase diagram as a guide to design optimum solution conditions for vapor diffusion experiments, and conversely how to understand the global shape of the protein phase diagram based on a few vapor diffusion experiments. These results extend the few studies that have attempted to use phase diagram approaches to improve protein crystallization.102,103,105 Conclusions Comparison of the effects of the protein additives ammonium sulfate, PEG 8000 and MPD shows that despite some differences in phase behavior among the additives, there is a general form of the phase diagram that is independent of the physical origin of protein interactions. For all three additives the aggregation line or liquid-liquid phase boundary has the same characteristic shape, and formation of aggregates, gels, and dense liquid phases is metastable to crystallization. This latter characteristic, which can be attributed to the short-range nature of protein interactions, is the fundamental reason that proteins crystallize. The present results therefore provide a common basis for understanding the phase behavior obtained with different additives, suggesting the universality of the concepts of phase behavior. The practical significance of the results can be seen in the use of the experimental phase diagram to optimize the solution conditions for hanging drop vapor diffusion experiments. The practical equivalence demonstrated between the information obtained from vapor diffusion and that from batch crystallization experiments makes it possible to understand protein phase behavior based on either method. Acknowledgment. We are grateful to the National Science Foundation for financial support (Grant Number BES-0519191).

References (1) George, A.; Wilson, W. W. Predicting protein crystallization from a dilute-solution property. Acta Crystallogr. Sect. D Biol. Crystallogr. 1994, 50, 361–365. (2) Tessier, P. M.; Lenhoff, A. M. Measurements of protein selfassociation as a guide to crystallization. Curr. Opin. Biotechnol. 2003, 14, 512–516. (3) Dumetz, A. C.; Snellinger-O’Brien, A. M.; Kaler, E. W.; Lenhoff, A. M. Patterns of protein-protein interactions in salt solutions and implications for protein crystallization. Protein Sci. 2007, 16, 1867– 1877. (4) George, A.; Chiang, Y.; Guo, B.; Arabshahi, A.; Cai, Z.; Wilson, W. W. Second virial coefficient as predictor in protein crystal growth. Methods Enzymol. 1997, 276, 100–110. (5) Piazza, R. Interactions in protein solutions near crystallization: A colloid physics approach. J. Cryst. Growth 1999, 196, 415–423. (6) Pan, X.; Glatz, C. E. Solvent effects on the second virial coefficient of subtilisin and solubility. Cryst. Growth Des. 2003, 3, 203–207.

Crystal Growth & Design, Vol. 9, No. 2, 2009 689 (7) Rosenbaum, D. F.; Zukoski, C. F. Protein interactions and crystallization. J. Cryst. Growth 1996, 169, 752–758. (8) Curtis, R. A.; Prausnitz, J. M.; Blanch, H. W. Protein-protein and protein-salt interactions in aqueous protein solutions containing concentrated electrolytes. Biotechnol. Bioeng. 1998, 57, 11–21. (9) Vivare`s, D.; Bonnete´, F. X-ray scattering studies of Aspergillus flavus urate oxidase: Towards a better understanding of PEG effects on the crystallization of large proteins. Acta Crystallogr. Sect. D Biol. Crystallogr. 2002, 58, 472–479. (10) Vivare`s, D.; Belloni, L.; Tardieu, A.; Bonnete´, F. Catching the PEGinduced attractive interaction between proteins. Eur. Phys. J. E 2002, 9, 15–25. (11) Kulkarni, A. M.; Chatterjee, A. P.; Schweizer, K. S.; Zukoski, C. F. Effects of polyethylene glycol on protein interactions. J. Chem. Phys. 2000, 113, 9863–9873. (12) Bloustine, J.; Virmani, T.; Thurston, G. M.; Fraden, S. Light scattering and phase behavior of lysozyme-poly(ethylene glycol) mixtures. Phys. ReV. Lett. 2006, 96, 87803–87807. (13) Berger, B. W.; Blamey, C. J.; Naik, U. P.; Bahnson, B. J.; Lenhoff, A. M. Roles of additives and precipitants in crystallization of calciumand integrin-binding protein. Cryst. Growth Des. 2005, 5, 1499–1507. (14) Tessier, P. M.; Johnson, H. R.; Pazhianur, R.; Berger, B. W.; Prentice, J. L.; Bahnson, B. J.; Sandler, S. I.; Lenhoff, A. M. Predictive crystallization of ribonuclease A via rapid screening of osmotic second virial coefficients. Proteins 2003, 50, 303–311. (15) Liu, W.; Bratko, D.; Prausnitz, J. M.; Blanch, H. W. Effect of alcohols on aqueous lysozyme-lysozyme interactions from static light-scattering measurements. Biophys. Chem. 2004, 107, 289–298. (16) Farnum, M.; Zukoski, C. Effect of glycerol on the interactions and solubility of bovine pancreatic trypsin inhibitor. Biophys. J. 1999, 76, 2716–2726. (17) Costenaro, L.; Zaccai, G.; Ebel, C. Understanding the crystallization of an acidic protein by dilution in the ternary NaCl-2-methyl-2,4pentanediol-H2O system. J. Cryst. Growth 2001, 232, 102–113. (18) Green, A. A.; Hughes, W. L. Protein fractionation on the basis of solubility in aqueous solutions of salts and organic solvents. Methods Enzymol. 1955, 1, 67–90. (19) Cohn, E. J.; Edsall, J. T. Proteins, Amino Acids and Peptides As Ions and Dipolar Ions; Reinhold Publishing Corporation: New York, NY, 1943. (20) McPherson, A. Crystallization of proteins from polyethylene glycol. J. Biol. Chem. 1976, 251, 6300–6303. (21) Polson, A.; Potgieter, G. M.; Largier, J. F.; Mears, G. E. F.; Joubert., F. J. The fractionation of protein mixtures by linear polymers of high molecular weight. Biochim. Biophys. Acta 1964, 82, 463–475. (22) Ray, W. J.; Bracker, C. E. Polyethylene-glycol: Catalytic effect on the crystallization of phosphoglucomutase at high salt concentration. J. Cryst. Growth 1986, 76, 562–576. (23) McPherson, A. Use of polyethylene glycol in the crystallization of macromolecules. Methods Enzymol. 1985, 114, 120–125. (24) Rupp, B. High-throughput crystallography at an affordable cost: The TB structural genomics consortium crystallization facility. Acc. Chem. Res. 2003, 36, 173–181. (25) Chayen, N. E.; Saridakis, E. Protein crystallization for genomics: Towards high-throughput optimization techniques. Acta Crystallogr. Sect. D Biol. Crystallogr. 2002, 58, 921–927. (26) Arakawa, T.; Timasheff, S. N. Mechanism of protein salting in and salting out by divalent cation salts: Balance between hydration and salt binding. Biochemistry 1984, 23, 5912–5923. (27) Arakawa, T.; Timasheff, S. N. Preferential interactions of proteins with salts in concentrated solutions. Biochemistry 1982, 21, 6545– 6552. (28) Kunz, W.; Henle, J.; Ninham, B. W. ’Zur Lehre von der Wirkung der Salze’ (about the science of the effect of salts): Franz Hofmeister’s historical papers. Curr. Opin. Colloid Interface Sci. 2004, 9, 19–37. (29) Hofmeister, F. Zur Lehre von der Wirkung der Salze. Arch. Exp. Pathol. Pharmakol. 1888, 24, 247–260. (30) McPherson, A. Introduction to protein crystallization. Methods 2004, 34, 254–265. (31) Arakawa, T.; Bhat, R.; Timasheff, S. N. Preferential interactions determine protein solubility in 3 component solutions: The MgCl2 system. Biochemistry 1990, 29, 1914–1923. (32) Grigsby, J. J.; Blanch, H. W.; Prausnitz, J. M. Diffusivities of lysozyme in aqueous MgCl2 solutions from dynamic light-scattering data: Effect of protein and salt concentrations. J. Phys. Chem. B 2000, 104, 3645–3650. (33) Rie`s-Kautt, M.; Ducruix, A. Crystallization of basic proteins by ion pairing. J. Cryst. Growth 1991, 110, 20–25.

690 Crystal Growth & Design, Vol. 9, No. 2, 2009 (34) Vaney, M. C.; Broutin, I.; Retailleau, P.; Douangamath, A.; Fafont, S.; Ducruix, A.; Ries-Kautt, M. Structural effects of monovalent anions on polymorphic lysozyme crystals. Acta Crystallogr. Sect. D Biol. Crystallogr. 2001, 57, 929–940. (35) Iyer, G. H.; Dasgupta, S.; Bell, J. A. Ionic strength and intermolecular contacts in protein crystals. J. Cryst. Growth 2000, 217, 429–440. (36) Petsev, D. N.; Thomas, B. R.; Yau, S.-T.; Vekilov, P. G. Interactions and aggregation of apoferritin molecules in solution: Effects of added electrolytes. Biophys. J. 2000, 78, 2060–2069. (37) Gilliland, G. L.; Davies, D. R. Protein crystallization: The growth of large-scale single crystals. Methods Enzymol. 1984, 104, 370– 381. (38) Gilliland, G. L. A biological macromolecule crystallization database: A basis for a crystallization strategy. J. Cryst. Growth 1988, 90, 51– 59. (39) McPherson, A.; Malkin, A. J.; Kuznetsov, Y. G. The Science of Macromolecular Crystallization. Structure 1995, 3, 759–768. (40) Arakawa, T.; Timasheff, S. N. Mechanism of poly(ethylene glycol) interaction with proteins. Biochemistry 1985, 24, 6756–6762. (41) Lee, J. C.; Lee, L. L. Interaction of calf brain tubulin with polyethylene glycols. Biophys. J. 1979, 25, A254. (42) Lee, J. C.; Lee, L. L. Preferential solvent interactions between proteins and polyethylene glycols. J. Biol. Chem. 1981, 256, 625–631. (43) Asakura, S.; Oosawa, F. Interaction between particles suspended in solutions of macromolecules. J. Polym. Sci. 1958, 33, 183–192. (44) Vrij, A. Polymers at interfaces and interactions in colloidal dispersions. Pure Appl. Chem. 1976, 48, 471–483. (45) Fuchs, M.; Schweizer, K. S. Macromolecular theory of solvation and structure in mixtures of colloids and polymers. Phys. ReV. E 2001, 64, 21514–21533. (46) Chatterjee, A. P.; Schweizer, K. S. Correlation effects in dilute particle-polymer mixtures. J. Chem. Phys. 1998, 109, 10477–10488. (47) Mahadevan, H.; Hall, C. K. Statistical-mechanical model of protein precipitation by nonionic polymer. AIChE J. 1990, 36, 1517–1528. (48) Dumetz, A. C.; Schweitzer, K. S.; Lenhoff, A. M.; Kaler, E. W. Protein interactions in polyethylene glycol solutions: An experimental study of depletion forces in the protein limit (in preparation). (49) Israelachvili, J. The different faces of poly(ethylene glycol). Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 8378–8379. (50) Furness, E. L.; Ross, A.; Davis, T. P.; King, G. C. A hydrophobic interaction site for lysozyme binding to polyethylene glycol and model contact lens polymers. Biomaterials 1998, 19, 1361–1369. (51) Sheth, S. R.; Leckband, D. Measurements of attractive forces between proteins and end-grafted poly(ethylene glycol) chains. Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 8399–8404. (52) Xia, J.; Dubin, P. L.; Kokufuta, E. Dynamic and electrophoretic light scattering of a water-soluble complex formed between pepsin and poly(ethylene glycol). Macromolecules 1993, 26, 6688–6690. (53) Schubert, P. F.; Finn, R. K. Alcohol precipitation of proteins: The relationship of denaturation and precipitation for catalase. Biotechnol. Bioeng. 1981, 23, 2569–2590. (54) Kirkwood, J. G. Proteins; Gordon and Breach: New York, NY, 1967. (55) Cohn, E. J. The Solubility of Proteins. In Proteins, Amino Acids and Peptides As Ions and Dipolar Ions; Cohn, E. J., Edsall, J. T., Eds.; Reinhold Publishing Corporation: New York, 1943. (56) Bonincontro, A.; De Francesco, A.; Matzeu, M.; Onori, M.; Santucci, A. Conformational changes of lysozyme in water-ethanol mixtures. Colloids Surf. B 1997, 10, 105–111. (57) Anand, K.; Pal, D.; Hilgenfeld, R. An overview on 2-methyl-2,4pentanediol in crystallization and in crystals of biological macromolecules. Acta Crystallogr. Sect. D Biol. Crystallogr. 2002, 58, 1722–1728. (58) Lehmann, M. S.; Mason, S. A.; McIntyre, G. J. Study of ethanollysozyme interactions using neutron diffraction. Biochemistry 1985, 24, 5862–5869. (59) Kamphuis, I. G.; Kalk, K. H.; Swarte, M. B. A.; Drenth, J. Structure of papain refined at 1.65 Å resolution. J. Mol. Biol. 1984, 179, 233– 256. (60) Teeter, M. M. Water structure of a hydrophobic protein at atomic resolution: Pentagon rings of water molecules in crystals of crambin. Proc. Natl. Acad. Sci. U. S. A. 1984, 81, 6014–6018. (61) Charron, C.; Kadri, A.; Robert, M. C.; Giegé, R.; Lorber, B. Structure of thaumatin crystallized in the presence of glycerol displaces water molecules in the structure of thaumatin. Acta Crystallogr. Sect. D Biol. Crystallogr. 2002, 58, 2060–2065. (62) Deshpande, A.; Nimsadkar, S.; Mande, S. C. Effects of alcohols on protein hydration: Crystallographic analysis of hen egg-white lysozyme in the presence of alcohols. Acta Crystallogr. Sect. D Biol. Crystallogr. 2005, 61, 1005–1008.

Dumetz et al. (63) Pittz, E. P.; Timasheff, S. N. Interaction of ribonuclease A with aqueous 2-mthyl-2,4-pentanediol. Biochemistry 1978, 17, 615–623. (64) Bolen, D. W. Effects of naturally occurring osmolytes on protein stability and solubility: Issues important in protein crystallization. Methods 2004, 34, 312–322. (65) Sousa, B. R. Use of glycerol, polyols and other protein stucture stabilizing agents in protein crystallization. Acta Crystallogr. Sect. D Biol. Crystallogr. 1995, 51, 271–277. (66) Taratuta, V. G.; Holschbach, A.; Thurston, G. M.; Blankschtein, D.; Benedek, G. B. Liquid-liquid phase separation of aqueous lysozyme solutions: Effects of pH and salt identity. J. Phys. Chem. 1990, 94, 2140–2144. (67) Muschol, M.; Rosenberger, F. Liquid-liquid phase separation in supersaturated lysozyme solutions and associated precipitate formation/crystallization. J. Chem. Phys. 1997, 107, 1953–1962. (68) Tanaka, S.; Yamamoto, M.; Ito, K.; Hayakawa, R. Relation between the phase separation and the crystallization in protein solutions. Phys. ReV. E 1997, 56, R67–R69. (69) Thomson, J. A.; Schurtenberger, P.; Thurston, G. M.; Benedek, G. B. Binary liquid phase separation and critical phenomena in protein/ water solution. Proc. Natl. Acad. Sci. U. S. A. 1987, 84, 7079–7083. (70) Berland, C. R.; Thurston, G. M.; Kondo, M.; Broide, M. L.; Pande, J.; Ogun, O.; Benedek, G. B. Solid-liquid phase boundaries of lens protein solution. Proc. Natl. Acad. Sci. U. S. A. 1992, 89, 1214– 1218. (71) Broide, M. L.; Berland, C. R.; Pande, J.; Ogun, O.; Benedek, G. B. Binary-liquid phase-separation of lens protein solutions. Proc. Natl. Acad. Sci. U. S. A. 1991, 88, 5660–5664. (72) Liu, C.; Lomakin, A.; Thurston, G. M.; Hayden, D.; Pande, A.; Pande, J.; Ogun, O.; Asherie, N.; Benedek, G. B. Phase separation in multicomponent aqueous-protein solutions. J. Phys. Chem. 1995, 99, 454–461. (73) Thurston, G. M. Liquid-liquid phase separation and static light scattering of concentrated ternary mixtures of bovine R and γB crystallins. J. Chem. Phys. 2006, 124, 134909–134919. (74) Lafont, S.; Veesler, S.; Astier, J.-P.; Boistelle, R. Comparison of solubilities and molecular interactions of BPTI molecules giving different polymorphs. J. Cryst. Growth 1997, 173, 132–140. (75) Grouzael, S.; Bonnete´, F.; Astier, J. P.; Ferte´, N.; Perez, J.; Veesler, S. Exploring bovine pancreatic trypsin inhibitor phase transitions. J. Phys. Chem. B 2006, 110, 19664–19670. (76) Grouazel, S.; Perez, J.; Astier, J.-P.; Bonnete´, F.; Veesler, S. BPTI liquid-liquid phase separation monitored by light scattering. Acta Crystallogr. Sect. D Biol. Crystallogr. 2002, 58, 1560–1563. (77) Vivare`s, D.; Veesler, S.; Astier, J.-P.; Bonnete´, F. Polymorphism of urate oxidase in PEG solutions. Cryst. Growth Des. 2006, 6, 287– 292. (78) Vivare`s, D.; Bonnete´, F. Liquid-liquid phase separations in urate oxidase/PEG mixtures: Characterization and implications for protein crystallization. J. Phys. Chem. B 2004, 108, 6498–6507. (79) Vivare`s, D.; Kaler, E. W.; Lenhoff, A. M. Quantitative imaging by confocal scanning fluorescence microscopy of protein crystallization via liquid-liquid phase separation. Acta Crystallogr. Sect. D Biol. Crystallogr. 2005, 61, 819–825. (80) Jion, A. I.; Goh, L.-T.; Oh, S. K. W. Crystallization of IgG1 by mapping its liquid-liquid phase separation curves. Biotechnol. Bioeng. 2006, 95, 911–918. (81) Wang, Y.; Annunziata, O. Comparison between protein-PEG interactions and the effect of PEG on protein-protein interactions using the liquid-liquid phase transition. J. Phys. Chem. B 2007, 111, 1222– 1230. (82) Annunziata, O.; Asherie, N.; Lomakin, A.; Pande, J.; Ogun, O.; Benedek, G. B. Effect of polyethylene glycol on the liquid-liquid phase transition in aqueous protein solutions. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 14165–14170. (83) Rosenbaum, D.; Zamora, P. C.; Zukoski, C. F. Phase behavior of small attractive colloidal particles. Phys. ReV. Lett. 1996, 76, 150– 153. (84) Zaccarelli, E.; Sciortino, F.; Tartaglia, P.; Foffi, G.; McCullagh, G. D.; Lawlor, A.; Dawson, K. A. Competition between crystallization and glassification for particles with short-ranged attraction. Possible applications to protein crystallization. Physica A 2002, 314, 539– 547. (85) Foffi, G.; McCullagh, G. D.; Lawlor, A.; Zaccarelli, E.; Dawson, K. A.; Sciortino, F.; Tartaglia, P.; Pini, D.; Stell, G. Phase equilibria and glass transition in colloidal systems with short-ranged attractive interactions: Application to protein crystallization. Phys. ReV. E 2002, 65, 31407.

Comparative Effects on Protein Phase Behavior (86) Anderson, V. J.; Lekkerkerker, H. N. W. Insights into phase transition kinetics from colloid science. Nature 2002, 416, 811–815. (87) Ilett, S. M.; Orrock, A. W. C.; Poon, K.; Pusey, P. N. Phase behavior of a model colloid-polymer mixture. Phys. ReV. E 1995, 51, 1344– 1352. (88) Gast, A. P.; Hall, C. K.; Russel, W. B. Polymer-induced phase separations in nonaqueous colloidal suspensions. J. Colloid Interface Sci. 1983, 96, 251–267. (89) Shah, S. A.; Chen, Y. L.; Schweizer, K. S.; Zukoski, C. Phase behavior and concentration fluctuations in suspensions of hard spheres and nearly ideal polymers. J. Chem. Phys. 2003, 118, 3350–3361. (90) Ramakrishnan, S.; Fuchs, M.; Schweizer, K. S.; Zukoski, C. Entropy driven phase transitions in colloid-polymer suspensions: Tests of depletion theories. J. Chem. Phys. 2002, 116, 2201–2212. (91) Dumetz, A. C.; Chockla, A. M.; Kaler, E. W.; Lenhoff, A. M. Protein phase behavior in aqueous solutions: Crystallization, liquid-liquid phase separations, gels and aggregates. Biophys. J. 2008, 94, 570– 583. (92) Cheng, Y.-C.; Lobo, R. F.; Sandler, S. I.; Lenhoff, A. M. Kinetics and equilibria of lysozyme precipitation and crystallization in concentrated ammonium sulfate solutions. Biotechnol. Bioeng. 2006, 94, 177–188. (93) Hopkins, F. G.; Pinkus, S. N. Observations on the crystallization of animal proteids. J. Physiol. 1898, 23, 130–136. (94) Judge, R. A.; Johns, M. R.; White, E. T. Protein purification by bulk crystallization: The recovery of ovalbumin. Biotechnol. Bioeng. 1995, 48, 316–323. (95) Sober, H. A., Ed. Handbook of Biochemistry. Selected Data for Molecular Biology; The Chemical Rubber Company: Cleveland, OH, 1970. (96) Bhat, R.; Timasheff, S. N. Steric exclusion is the principal source of the preferential hydration of proteins in the presence of polyethylene glycols. Protein Sci. 1992, 1, 1133–1143.

Crystal Growth & Design, Vol. 9, No. 2, 2009 691 (97) King, M. V.; Magdoff, B. S.; Adelman, M. B.; Harker, D. Crystalline forms of bovine pancreatic ribonuclease: Techniques of preparation, unit cells, and space groups. Acta Crystallogr. Sect. D Biol. Crystallogr. 1956, 9, 460. (98) King, M. V.; Bello, J.; Pignataro, E. H.; Harker, D. Crystalline forms of bovine pancreatic ribonuclease. Some new modifications. Acta Crystallogr. 1962, 15, 144–147. (99) ten Wolde, P. R.; Frenkel, D. Enhancement of protein crystal nucleation by critical density fluctuations. Science 1997, 277, 1975– 1978. (100) ten Wolde, P. R.; Frenkel, D. Enhanced protein crystallization around the metastable critical point. Theor. Chem. Acc. 1999, 101, 205– 208. (101) de Gennes, P.-G. Suspensions colloidales dans un me´lange binaire critique. C. R. Acad. Sci., Ser. IIb: Mec., Phys., Chim., Astron. 1981, 292, 701–704. (102) Anderson, M. J.; Hansen, C. L.; Quake, S. R. Phase knowledge enables rational screens for protein crystallization. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 16746–16751. (103) Saridakis, E.; Chayen, N. E. Systematic improvement of protein crystals by determining the supersolubility curves of phase diagrams. Biophys. J. 2003, 84, 1218–1222. (104) McPherson, A. Crystallization of Biological Macromolecules; Cold Spring Harbor Laboratory Press: New York, 1999. (105) Saijo, S.; Sato, T.; Tanaka, N.; Ichiyanagi, A.; Sugano, Y.; Shoda, M. Precipitation diagram and optimization of crystallization conditions at low ionic stength for deglycosylated dye-decolorizing peroxidase from a basidiomycete. Acta Crystallogr. Sect. D Biol. Crystallogr. 2005, 61, 729–732.

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