CRYSTAL GROWTH & DESIGN
Simultaneous Monitoring of Protein and (NH4)2SO4 Concentrations in Aprotinin Hanging-Drop Crystallization Using Raman Spectroscopy
2002 VOL. 2, NO. 6 511-514
Rosana E. Tamagawa,† Everson A. Miranda,† and Kris A. Berglund*,‡,§ Department of Bioprocesses, School of Chemical Engineering, Campinas State University, Campinas, SP, Brazil, Departments of Chemistry, Chemical Engineering & Materials Science, and Agricultural Engineering, Michigan State University, East Lansing, Michigan 48824, and Division of Chemical Engineering Design, Luleå University of Technology, SE-971 87 Luleå, Sweden Received June 23, 2002
ABSTRACT: In the present study the use of fiber optic Raman spectroscopy for the in situ monitoring of aprotinin supersaturation in a hanging-drop crystallization is described. The crystallizing agent, (NH4)2SO4, is Raman active, which allows monitoring of the salt concentration in the drop during the whole hanging-drop crystallization process in addition to monitoring of the aprotinin concentration. Through the continuous measuring of protein and salt concentrations, supersaturation in the drop solution was measured in real time. Moreover, through the monitoring of protein and salt concentrations, the cocrystallization of aprotinin and (NH4)2SO4 was observed. Introduction The key to control of crystallization is the ability to monitor and control the supersaturation. Previously, a number of methods were attempted for monitoring, including dynamic light scattering,1,3 small-angle X-ray scattering,2,5 differential calorimetry,4 and time-resolved fluorescence.10 These studies achieved different degrees of success in monitoring the protein concentration and supersaturation, but none included the simultaneous measurement of the concentration of precipitating agent (usually salt). Schwartz and Berglund11 demonstrated the use of Raman spectroscopy for the real-time monitoring of lysozyme supersaturation in hanging-drop crystallization. Recently the monitoring of aprotinin (a serine protease inhibitor with pharmaceutical applications composed of a single chain of 58 amino acid residues; 6511 Da) supersaturation in the presence of NaCl using Raman spectroscopy was demonstrated and the supersaturation was controlled to increase the product aprotinin crystal size.12 In the present work, the approach of monitoring protein supersaturation with Raman spectroscopy is extended to monitor salt ((NH4)2SO4) concentration in addition to protein concentration during crystallization in the hanging drop. This approach provides a more realistic determination of supersaturation, since solubility also depends on the salt concentration, a consideration not included in previous studies. Materials Solutions used in the crystallization experiments were prepared with deionized water and were filtered through 0.45 µm pore size filters. Aprotinin was obtained from Sigma (USA), and all reagents were of analytical grade. Siliconized glass cover slides used for hanging drop crystallization were pur* To whom correspondence should be addressed at the Luleå University of Technology. † Campinas State University. ‡ Michigan State University. § Luleå University of Technology.
chased from Hampton Research, Laguna Niguel, CA. IZIT, an organic reagent by Hampton Research, was used to distinguish between salt and protein crystals. Raman spectra were collected with a Kaiser Optical Systems, Inc. HoloLab Series 5000.
Methods Acquisition and Calibration of Raman Spectra. Raman spectra (Figure 1) were collected using a HoloLab Series 5000 from Kaiser Optical Systems (USA), which employs a 100 mW external cavity stabilized diode laser at 785 nm for sample illumination. A CCD camera, a spectrograph, and a fiber optic probe complete this system. Remote sampling was accomplished by employing a fiber optic probe attached to a 10× microscope objective to focus the incident beam. The calibration model was based on spectra of 20 standard solutions ranging in concentration from 0 to 100 mg/mL of protein and from 0.0 to 2.0 M of (NH4)2SO4. Solutions were prepared in 50 mM sodium acetate buffer, pH 4.5. All spectra were acquired from approximately 10 µL volume and were sums of 10 accumulations collected over 5 s, each at 8 cm-1 resolution. A partial least-squares (PLS) regression model was built using QuantIR multivariate regression analysis software by ASIMettler Toledo (USA). Hanging-Drop Crystallization. Ten-microliter droplets of solution containing protein and salt were placed on glass cover slides which were inverted and suspended over vessels with 10 mL of the reservoir solution. Reservoir solutions contained higher concentrations of salt; therefore, water is transferred from the drop to the reservoir in order to equilibrate the vapor pressure between the two solutions. The top of the vessels was treated with silicone in order to ensure an airtight seal. To ensure that the resulting crystals were composed of protein, 1 µL of an organic dye, IZIT, was added to drops containing the crystals. Protein crystals contain high contents of solvent distributed through microscopic channels, which allow the organic reagent to permeate through the structure, giving the crystal a blue color. On the other hand, salt crystals, being more compact, do not absorb the reagent and remain transparent.
Results and Discussion Calibration of Raman Spectra. Raman spectra of standard solutions were collected as described and
10.1021/cg025544m CCC: $22.00 © 2002 American Chemical Society Published on Web 08/24/2002
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Figure 1. Raman spectrum of an aqueous aprotinin-(NH4)2SO4 solution at a protein concentration of 100 mg/mL and (NH4)2SO4 concentration of 1.0 M in 50 mM sodium acetate buffer at pH 4.5 and a temperature of 24 °C. Laser illumination was 100 mW at 785 nm.
Figure 2. Aprotinin solubility at 24 °C and pH 4.5 in the presence of (NH4)2SO4 measured using Raman spectroscopy. Solutions were buffered with 50 mM sodium acetate. Volumes of the drops and reservoir were 10 µL and 10 mL, respectively. Drops initially contained 70, 40, 20, and 10 mg/mL of aprotinin and 1.0 M of (NH4)2SO4, and the respective reservoirs contained 1.5, 1.6, 1.7, and 1.8 M of (NH4)2SO4.
treated with a PLS calibration using QuantIR. The peak regions selected to correlate spectra and solution composition were 410-535, 825-880, 924-955, 1204-1230, 2880-2980, 1025-1050, and 1178-1080 cm-1. These regions correspond to aprotinin and (NH4)2SO4 vibrational bands. Remaining regions of the spectra containing solvent and glass scattering as well as overlapped peaks resulted in a poor calibration and were excluded from the model. The standard error determined by a leave-one-out cross-validation was 0.54 mg/mL of aprotinin and 0.03 M of salt. Solubility Determination. Aprotinin solubility in the presence of (NH4)2SO4 (Figure 2) was determined by measuring the equilibrium concentration in hangingdrop crystallizations. Droplets (10 µL) of aprotinin solution were suspended above vessels with 10 mL of the respective crystallizing solution. Drops contained initially 70, 40, 20, and 10 mg/mL of aprotinin and 1.0 M of (NH4)2SO4, and the respective reservoirs contained 1.5, 1.6, 1.7, and 1.8 M of (NH4)2SO4. The dropreservoir systems were left at room temperature for 3 weeks to allow equilibrium. Raman spectra from the liquid phase were collected to determine protein equi-
librium concentration. The experiment was conducted in triplicate, and the error bars (standard deviation) in Figure 2 reflect the experimental reproducibility. As reported by Lafont et al.,7 aprotinin solubility decreases with an increase in (NH4)2SO4 concentration. The same behavior was observed in aprotinin crystallization in the presence of NaCl.8,12 Monitoring of Supersaturation in the HangingDrop Crystallization. In a typical experiment, a 10 µL drop of solution containing 30 mg/mL of aprotinin and 1.26 M (NH4)2SO4 was suspended above a reservoir containing 2.2 M aqueous (NH4)2SO4 (both the drop and reservoir solutions were buffered with 50 mM sodium acetate to pH 4.5). The Raman fiber optic probe was then placed over the drop-reservoir system, and with the laser beam focused in the drop solution, spectra from the drop solution were collected continuously at 10 s intervals. Using the spectral differences between the solution and solid phases, it was possible to ensure that the liquid and not the solid phase was being evaluated. The spectrum of the solid phase results in higher intensities with narrower half-widths and additional bands as compared to the spectrum of the liquid phase. Therefore, if a crystal grew in the path of the incident beam, the spectral intensities increased abruptly. Upon observation of a crystal in the incident beam, the reservoir and drop system was adjusted in order to bring the laser focus from the solid to the liquid phase. Through this procedure, it was ensured that the solution-phase aprotinin and (NH4)2SO4 concentrations were monitored during the hanging-drop crystallization process as shown in Figure 3. By this monitoring it was possible to distinguish when the nucleation stage occurred and the time when equilibrium was achieved. The increase in aprotinin and salt concentrations in the first 500 min (Figure 3) corresponds to evaporation leading to nucleation, while the gradual decrease in the next 500 min represents primarily crystal growth. The concentration decrease is due to the transport of the solute from the liquid to the solid phase, noting that only the solution phase is being monitored. Equilibrium (solubility) was attained when the concentration reached a constant value. In addition to distinguishing between crystallization steps, the crystallization could be monitored in terms
Aprotinin Hanging-Drop Crystallization
Figure 3. Solution aprotinin and (NH4)2SO4 concentration changes with time in hanging-drop crystallization. Initially, the drop volume was 10 µL, the reservoir volume was 10 mL, and the reservoir contained 2.2 M (NH4)2SO4. The temperature was 24 °C, and the solutions were buffered at pH 4.5 with 50 mM sodium acetate.
Figure 4. Aprotinin supersaturation monitoring during a hanging-drop crystallization. Initially, the drop volume was 10 µL, the reservoir volume was 10 mL, and the reservoir contained 2.2 M (NH4)2SO4. The temperature was 24 °C, and the solutions were buffered at pH 4.5 with 50 mM sodium acetate.
of the solubility diagram (Figure 4). In this diagram, protein and salt concentrations in the drop measured during the initial concentration stage were plotted together with the solubility curve (solid line). Due to transport from the droplet to the reservoir, protein and salt concentrations increased gradually, leading to the supersaturation of the solution with respect to aprotinin. When the metastable limit for supersaturation was reached, nucleation started and consequently the protein concentration decreased (dashed arrow) until the solution reached the equilibrium (solubility curve). Only protein concentration is presented as a decreasing variable in the diagram, even though salt concentration also decreased during crystal growth (Figure 3). Figure 4 depicts the salt concentration in the drop at which aprotinin (target molecule) crystallized. The cocrystallization of aprotinin and (NH4)2SO4 was unexpected and is discussed below. Cocrystallization of Aprotinin and Ammonium Sulfate. As shown in Figure 3, during the crystallization of aprotinin in the presence of (NH4)2SO4, the solution-phase (NH4)2SO4 concentration decreased in parallel to the aprotinin concentration decrease. This
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Figure 5. Solution aprotinin concentration as a function of (NH4)2SO4 concentration in drops A and B during the nucleation stage. Drops A and B initially contained 50 and 30 mg/ mL of aprotinin, respectively. Additionally, drops A and drop B contained approximately 1.2 M (NH4)2SO4 and were suspended above reservoirs containing 2.2 M of the same salt. The temperature was 24 °C.
behavior was initially associated with the formation of separate aprotinin and (NH4)2SO4 crystals. This hypothesis was consistent with the presence of rod-shaped crystals similar in shape to (NH4)2SO4 crystals. However, after the crystal composition was checked, employing IZIT, as described, this hypothesis was discarded. With the addition of the dye reagent to the drop, all crystals turned blue, indicating that they were aprotinin crystals. Therefore, the remaining possibility was the cocrystallization of protein and salt in a combined form. In fact, it is known that the structure of protein crystals may include the presence of some cations and anions. Exposed hydrophilic groups on these macromolecules can bind not only water molecules but also a variety of ions such as Ca2+, Na+, Cl-, and SO42-, particularly under crystallization conditions where they are found at high concentrations.9 Aprotinin and (NH4)2SO4 cocrystallization is consistent with the work of Hamiaux et al.6 During the refinement of the aprotinin crystal form obtained in the presence of (NH4)2SO4, Hamiaux et al. found the presence of sulfate ions in the crystal structure. They also noticed that the increase of (NH4)2SO4 concentration in undersaturated and supersaturated solutions of aprotinin resulted in oligomerization in a decameric form. Since the crystal structure solved by X-ray diffraction shows a compact decamer, they proposed that, in supersaturated solution, aprotinin crystal growth proceeds by the decamer stacking. Within a decamer five strong diffraction peaks were found and interpreted as sulfate ions located at the pentamer-pentamer location. The ratio of aprotinin and (NH4)2SO4 participating in the crystal packing was estimated by measuring aprotinin and (NH4)2SO4 concentrations throughout the crystallization process. Two drops, A and B, containing 1.2 M of (NH4)2SO4 and 50 and 30 mg/mL of aprotinin, respectively, were suspended above reservoirs containing 2.2 M of (NH4)2SO4. Plots of aprotinin concentration versus salt concentration for the nucleation and crystal growth steps were prepared and are shown in Figures 5 and 6. Drop A, which had a higher initial protein concentration than drop B, maintained a higher solution protein
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Conclusions Raman spectroscopy was demonstrated for simultaneous monitoring of aprotinin and (NH4)2SO4 concentrations during hanging-drop crystallization. This measurement allowed an accurate determination of supersaturation, which is also dependent on the salt concentration. The monitoring of aprotinin crystallization in the presence of (NH4)2SO4 showed the depletion of both components from the liquid phase, indicating the cocrystallization of protein and salt. Raman spectroscopy has unique sampling attributes for protein crystallization studies and allows measurements not possible with other techniques. Figure 6. Protein and salt depletion from solution during the crystal growth stage of the cocrystallization of aprotinin and (NH4)2SO4. Initial protein concentrations were 50 mg/mL of aprotinin (drop A) and 30 mg/mL of aprotinin (drop B). The (NH4)2SO4 concentration was 1.2 M in the drop and 2.2 M in the reservoir. The temperature was 24 °C. The lines represent least-squares fits to the data.
concentration throughout the nucleation stage and consequently reached a higher supersaturation (Figure 4). In the crystal growth stage (Figure 6), the decrease in protein concentration was linearly proportional to the decrease in salt concentration for both drops. The slopes of the lines, however, were different for each drop (6.5 × 10-4 [M aprotinin/M (NH4)2SO4] for drop A and 3.3 × 10-4 [M aprotinin/M (NH4)2SO4] for drop B). This indicates that the ratio of protein to salt depletion from solution depends on the ratio of protein and salt concentration when crystallization started, where the higher the aprotinin to (NH4)2SO4 ratio in the drop when crystallization started, the higher the ratio in the crystal. The reciprocals of the linear coefficients provide the amount of salt present in the solid phase for an amount of protein, resulting in 162 and 249 mol of salt per mol of aprotinin for crystals A and B, respectively. These values are 1 order of magnitude higher than the value determined by Hamiaux et al.6 Since IZIT does not detect salt crystals, it is possible that some of the (NH4)2SO4 may have have crystallized out of solution and formed crystals that remained undetected. The stoichiometry of the crystallization merits further study beyond this initial feasibility study.
Acknowledgment. We wish to thank the CAPES and FAPESP, Brazil, for financial support of R.E.T. Additional support from the Center for New Plant Products and Processes at Michigan State University is also appreciated. Kaiser Optical Systems, Inc., is also thanked for supplying the Raman spectrometer used in this study. References (1) Ansari, R. R.; Suh, K. I.; Arabshahi, A.; Wilson, W. W.; Bray, T. L.; DeLucas, L. J. J. Cryst. Growth 1996, 168, 216-226. (2) Bonnete´, F.; Vidal, O.; Robert, M. C.; Tardieu, A. J. Cryst. Growth 1996, 168, 185-191. (3) Boyer, M.; Roy, M.-O.; Jullien, M. J. Cryst. Growth 1996, 167, 212-220. (4) Darcy, P. A.; Wiencek, J. M. J. Cryst. Growth 1999, 196, 243-249. (5) Ducruix, A.; Guilloteau, J. P.; Rie`s-Kaut, M.; Tardieu, A. J. Cryst. Growth 1996, 168, 28-29. (6) Hamiaux, C.; Pe´rez, J.; Prange´, T.; Veesler, S.; Rie`s-Kautt; Vachette, P. J. Mol. Biol. 2000, 297, 697-712. (7) Lafont, S.; Veesler, S.; Astier, J. P.; Boistelle, R. J. Cryst. Growth 1997, 173, 132-140. (8) Lafont, S.; Veesler, S.; Astier, J. P.; Boistelle, R. J. Cryst. Growth 1994, 143, 249-255. (9) McPherson, A. Crystallization of Biological Macromolecules; Cold Spring Harbor Laboratory Press: New York, 1999. (10) Pan, B.; Berglund, K. A. J. Cryst. Growth 1997, 171, 226235. (11) Schwartz, A.; Berglund, K. A. J. Cryst. Growth 1999, 203, 599-603. (12) Tamagawa, R. E.; Miranda, E. A.; Berglund, K. A. Cryst. Growth Des, in press.
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