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* Department of Anatomy and Structural Biology, Albert Einstein College of Medicine and Montefiore Hospital, Comprehensive Sickle Cell Center, The Bronx, New York; Department of Chemical Engineering, University of Houston, Houston, Texas; Department of Physics, Hunter College of the City University of New York, New York; and ¶ Department of Medicine (Division of Hematology) and Department of Physiology and Biophysics, Albert Einstein College of Medicine and Montefiore Hospital, Comprehensive Sickle Cell Center, The Bronx, New York
Correspondence: Address reprint requests to Peter G. Vekilov, Dept. of Chemical Engineering, University of Houston, Houston, TX 77204-4004. Tel.: 713-743-4315; Fax: 713-743-4323; E-mail: vekilov{at}uh.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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Lys) (Hunt and Ingram, 1958
). In its oxygenated, R state, HbC forms crystals inside red blood cells. The intraerythrocytic crystals contribute to the clinical pathogenesis of the CC disease, a mild and chronic hemolytic anemia (Lawrence et al., 1991; Lessin et al., 1969). Oxy-HbC crystals also form in red cells of patients with the SC disease, who are doubly heterozygous for both HbS and HbC. SC disease is a severe, life-threatening condition (Nagel and Lawrence, 1991). Thus, data on the mechanisms and dynamics of HbC crystallization, including at the molecular level, contributes to the understanding of the pathogenesis of the CC and SC diseases, and may be relevant to the other protein condensation diseases: sickle cell anemia (Eaton and Hofrichter, 1990), the eye cataract (Benedek et al., 1999), Alzheimer's (Kelly, 2000), the prion diseases (Bucciantini et al., 2002; Koo et al., 1999), and others.
Previous observations of the formation of HbC crystals in osmotically dehydrated red blood cells were interpreted in terms of a pathway involving formation of small ribbon-like "paracrystals," followed by their alignment into tetragonal and hexagonal crystals (Charache et al., 1967). The suggested mechanism of alignment of the microribbons is similar to the one implied for the other ß6 Hb variant, deoxy-HbS (Bluemke et al., 1988; Lessin et al., 1969; Makinen and Sigountos, 1984; Potel et al., 1984). This pathway is at odds with the crystallization mechanism established for a broad variety of proteins under a broad range of conditions (McPherson et al., 2000; Vekilov and Chernov, 2002)—in all cases crystallization occurs by the association of single molecules into an existing crystal. Thus, in situ monitoring of the elementary acts of molecular attachment to the crystals are needed to decide if indeed hemoglobin crystallization follows a pathway distinct from the one of most other proteins.
In preceding studies, we found that, similar to many other proteins, the solubility of the protein has a strong retrograde dependence on temperature (Feeling-Taylor et al., 1999). We analyzed the thermodynamics of HbC crystallization and found a high positive enthalpy (Vekilov et al., 2002). This means that the specific interactions favor HbC molecules in the solute state, and HbC crystallization is only possible because of a huge entropy gain, likely stemming from the release of up to 10 water molecules per protein intermolecular contact. Such contacts have been defined as hydrophobic attraction (Chandler, 2002; Eisenberg and Kauzmann, 1969). Thus, our data suggest that the greater propensity for R-state HbC crystallization is attributable to the increased hydrophobicity resulting from conformational changes that accompany the HbC mutation. We also found that the interactions between the HbC, HbA, and HbS molecules in solution are dominated by chemically specific binding between the surfaces of Hb molecules that are adjacent in the respective solid phase, and that the contributions of the electrostatic interactions are insignificant at electrolyte concentrations 
0.05 M (Vekilov et al., 2002). These observations do not reveal any specificity of HbC crystallization that may underlie a crystallization mechanism different from the one established for many other proteins. In addition, microscopic observations and determinations of the nucleation statistics showed that the crystals of HbC nucleate and grow by the attachment of native molecules from the solution (Vekilov et al., 2002).
This article is focused on the mechanisms of growth of the HbC crystals. We employ atomic force microscopy (AFM) in situ, during the growth of the crystals from supersaturated solutions, or in saturated solutions, i.e., when the crystals are in a dynamic equilibrium with the solution. Because of the higher stability of CO-HbC compared with Oxy-HbC, we used the carbomonoxy form as representative of the R-state HbC. The structure of carbomonoxy-HbC was recently determined (Dewan et al., 2002) using crystals grown under conditions analogous to those employed here.
Crystallization was carried out at concentration of the potassium phosphate buffer of 1.9 M, at pH 7.35. These high concentrations of the electrolyte significantly lower the solubility of the hemoglobin and allow high relative supersaturations at low concentrations of the precious protein. Attempts to carry out crystallization studies at low buffer concentrations yielded crystals with phosphate in the range 0.6–1.6 M only if polyethylene glycol is added; at even lower phosphate concentrations no crystallization of CO-HbC occurs. The interactions and thermodynamics in protein-water-polymer systems are completely different from a system that does not contain the polymer and we did not consider results obtained with PEG relevant to the goals of this investigation. Obviously, the phosphate concentration used here is significantly higher than the one in the red blood cells. Our failure to obtain crystals at low phosphate concentrations suggests that in vivo crystallization of R-state HbC is a complex phenomenon that depends on many factors. We view the investigations reported here as a first step in the understanding of these processes.
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EXPERIMENTAL PROCEDURES |
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.
Growth of crystals on glass substrates for AFM observations
Coverslips treated with trimethylchlorosilane were glued to iron disks and used as glass substrates for AFM crystal growth studies. 20 µl aliquots of 20 mg mL–1 CO-HbC in 1.9 M KH2PO4/K2HPO4 at pH 7.35 were dispensed on each disk. To avoid evaporation, the droplets were covered with glass covers, hermetically sealed and kept for a few hours in a controlled temperature chamber at 
22°C. Typically, this led to the formation of 3–20 crystals of sizes ranging from 20 to 200 µm firmly attached to the glass bottom. The crystals always had the tetragonal bipyramid habit, consisting of eight {101} faces, typical of the tetragonal crystals of R-state HbC (see, e.g., Hirsch et al., 1988). Droplets with three to five crystals were selected and the crystals were washed by 1.9M KH2PO4/K2HPO4, pH 7.35. The iron disks with the glass covers holding the crystals were then magnetically mounted on the AFM scanner. The fluid AFM cell was filled with the crystallizing solution and imaging commenced.
Atomic force microscopy observations
We used a multimode atomic force microscope Nanoscope IIIa (Digital Instruments, Santa Barbara, CA). All images were collected in situ during the growth of the crystals using the less intrusive tapping mode. This mode allows visualization of adsorbed protein and impurity species—tip impact in contact imaging mode often prevents such imaging. We used the standard SiN tips. The tapping drive frequency was in the range of 25–31 kHz, within the range used by other authors (Yip and Ward, 1996), adjusted to the resonance value of the used tip. Other scanning parameters were adjusted such that continuous imaging affected neither the surface structure nor the process dynamics. For verification, we varied the scan sizes and the time elapsed between image collections, and saw that neither the spatial nor the temporal characteristics of the processes changed. For details and tests about the determination of the maximum resolution of 16 Å, and the calibration of the AFM imaging technique with other studied proteins, see Yau et al. (2000a). For calibration of the AFM technique with HbC crystals, we note that the experiments with HbC crystals in supersaturated solutions (see below) revealed that the thickness of a layer on a (101) face is 55 Å, and the periodicity along the [
01]-axis is 200 Å, in agreement with the x-ray crystal structure, according to which the respective values should be 53 and 202 Å (Dewan et al., 2002).
The characteristic shape of the crystals with the pyramid apex in the [001] (or c) direction allowed an easy correlation between the crystallographic directions and the AFM image orientation.
Temperature control and solubility
Temperature in the laboratory was stabilized to 22°C. No additional temperature control of the solution in the AFM fluid cell was employed. Insertion of a thermocouple in the crystallizing solution revealed that its temperature was 27 ± 0.7°C.
The solubility of CO-HbC in the solution with the used composition at the temperature of the crystallization, 27°C, was determined by monitoring trains of growth steps (see below) and gradually decreasing the CO-HbC concentration in the solution. We found that at C = 1.2 ± 0.3 mg mL–1, the propagation of the steps stopped, and when the concentration was lowered below this value, step movement was backward and the crystal was dissolving. We assign the width of the solubility range to the high sensitivity of the solubility to the concentration of the phosphate buffer: slight variations in the buffer concentration between preparations result in noticeable solubility variations. This value was chosen as the CO-HbC solubility at the temperature of the experiments. It agrees with the value extrapolated from the solubility in the 10–16°C range by using the crystallization enthalpy of 155 kJ mol–1 (Vekilov et al., 2002).
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RESULTS |
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). Indeed, the 41 axis oriented along the [001] direction, i.e., at an angle with the observed (101) face and perpendicular to the [010] direction, rotates each of the four molecules by 90° with respect to its neighbors in the quartet, exposing different sides of the HbC molecule.
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. These figures reveal that if the crystal surface was fully determined by the bulk structure, neither the grooves nor the four member strands should be present, i.e., the surface is reconstructed. The surface reconstruction is discussed in detail below.
The growth mechanisms: generation of new crystal layers
Fig. 2 a and many other similar images reveal that the crystals grow by a two step mechanism: 1), new layers are generated by a surface nucleation process; and 2), these layers incorporate building blocks from the solution and spread to cover the whole facet. Note that the generation of a subsequent layer occurs while the underlying layer is still growing. This leads to many layers spreading and chasing and meeting one another on the crystal surface. This crystallization mechanism has been postulated by Volmer in the 1930s (Volmer, 1939) and observed for numerous small-molecule, protein and virus crystals (Giesen et al., 1996; Malkin et al., 1996; McPherson et al., 2000; Yau et al., 2000a; Yip and Ward, 1996).
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; Gratz et al., 1993; Teng et al., 1998), and has been observed with proteins (Durbin and Carlson, 1992; Durbin and Feher, 1996; Land et al., 1997; Malkin et al., 1996; Reviakine et al., 2003). Note the hollow core around the dislocation axis. The core lowers the strain due to the misfit of the lattice layers at the screw dislocation. Its width is determined by the balance of this strain with the surface free energy between the solution and the crystal. A consistent thermodynamic theory reveals that the core width also depends on the elastic properties of the crystal and the supersaturation (Van den Hoek et al., 1982; Vekilov et al., 1990). Hollow cores around screw dislocation have been seen for many inorganic (De Yoreo et al., 1997) and a protein crystal (Kuznetsov et al., 1999). The significance of the observation of hollow cores with protein crystals is that it indicates that the elastic properties of the protein crystals and their response to lattice strain are not principally different from those of small-molecule crystals (Chernov, 2003; Vekilov and Chernov, 2002).
The growth mechanisms: incorporation of molecules into steps
Zooming in on the edge of the growing layer, Fig. 3, we find that the layer thickness is 55 Å—close to the prediction of 53 Å based on the x-ray structure (Dewan et al., 2002). Furthermore, we see that the molecular rows along the a (or b) axes are in perfect alignment with the underlying layers, as one would expect for the tetragonal lattice of the CO-HbC crystals. The edge of the unfinished layers is rough, and the characteristic lengthscale of the roughness equals one molecular dimension. This is only possible if molecules join the crystal one by one. We conclude that the building blocks of CO-HbC crystals are single protein molecules.
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], [010] , and [
] crystallographic directions computed from the respective displacements of steps between pairs of images in Fig. 4. We see that the velocities are not steady and randomly change in time because of the intrinsic stochasticity of the molecular incorporation processes, i.e., the number of molecules joining the step for a certain time is a random variable over the times of imaging in Fig. 4. Still, if we average the data for each direction in Fig. 5, we get roughly equal averages, in agreement with the isotropy of step propagation discussed above.
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= 1.1 nm s–1. We can use this value for a rough evaluation of the step kinetic coefficient ß, a parameter characterizing the time-averaged kinetics of incorporation of the HbC molecules into crystals. We use the definition of ß (Chernov, 1984)
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= 
(54 x 195 x 195) Å3 = 5.1 x 10–19 cm3 (Dewan et al., 2002) is the volume that a molecule occupies in the crystal. We get ß 
2 x 10–5 cm s–1, at the lower end of the range of ß for other proteins, 10–2 – 10–5 cm s–1 (Reviakine et al., 2003).
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).
Impurities and crystal growth
Dynamic light scattering investigations of crystallizing CO-HbC solutions revealed that aggregates of HbC form in solutions of concentrations higher than a few mg mL–1; a typical distribution of the protein species is shown in Fig. 7 a (for details about the technique, see Vekilov et al., 2002). In contrast to the distribution in low-concentration HbC solutions, which show a monodisperse sample with a characteristic molecular size of 5.5 nm (Vekilov et al., 2002), the distribution in Fig. 7 a shows the presence of at least two populations. The smaller-size population has an apparent mean diameter of 9 nm and these are likely the native HbC molecules. The increase from the typical size of the HbC molecules of 5.5 nm is attributable to the intermolecular attraction in the high-phosphate buffer solution that leads to lower diffusion coefficient (Schmitz, 1990). The lower diffusivity is interpreted by the data-processing algorithms as a size increase (Provencher, 1979, 1982a,b). The larger size population has a mean size of 30 nm and these are likely aggregates of HbC molecules. AFM monitoring of the growth interface revealed that the aggregates are strongly adsorbed at the growth interface, where they may serve as centers for the nucleation of new crystal layers. Fig. 7 b shows that the adsorbed aggregates often pin down the growth layers and slow down their propagation. Cross-section profiles of the images of the aggregates in Fig. 7 b and in other images with higher magnification reveal that their vertical dimensions are in the range 10–40 nm (the lateral dimensions may be misleading because of tip-sample convolution effects). This roughly agrees with the 30-nm aggregates detected by the light scattering determination in Fig. 7 a.
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DISCUSSION |
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01] direction. None of the produced structures possess the essential feature of Fig. 1 a—four-member strands separated by grooves. Thus, we conclude that the surface layer of the crystal, as seen in Fig. 1 a and other similar images is "reconstructed"; numerous instances of reconstruction of the surface layers of inorganic crystals have been documented (Swartzentruber, 1998; Williams and Bartelt, 1991).
The four-molecule-wide strands on the surfaces of the CO-HbC crystals are akin to the two-member strands observed by electron microscopy on the surface of HbS crystals grown from solutions containing PEG (Bluemke et al., 1988; Lessin et al., 1969; Makinen and Sigountos, 1984; Potel et al., 1984). In these studies, the presence of such strands was the main piece of evidence supporting the conclusion of a growth mechanism of HbS crystal involving alignment of the preformed fibers in the solution. Since fibers, ribbons, and other linear objects have been observed in crystallizing solutions of CO-HbC (Hirsch et al., 2001), one may speculate that the observations of the strands support the fiber-alignment mechanism of formation of HbC crystals. However, the results in Figs. 3 and 4 above unambiguously show that the growth of crystals occurs by the attachment of single molecules. In another article (Vekilov et al., 2002), we presented compelling evidence that the nucleation of the CO-HbC crystal is also a process of self-assembly of single molecules and the microribbons and other noncrystalline phases present in the solution only play auxiliary roles in the crystallization process. The overall conclusion emerging from these investigations is that the crystallization mechanisms of HbC, in concentrated phosphate buffer, are by no means different from those found for other proteins (see, e.g., McPherson, 1999; McPherson et al., 2000; Yau et al., 2000a).
Impurities and crystallization
The observations of the impurity effects on the growth of HbC crystals provide some additional insight into a general issue of protein crystallization—that of the nature and role of impurity species. It is commonly thought that crystallization is a method for protein purification. However, there have been numerous crystal growth studies wherein crystal imperfections arise from the incorporation of "impurities," e.g., (Vekilov, 2003). It was found that "impurities," i.e., microheterogeneous molecules of the crystallizing protein, (e.g., aggregates and clusters of this protein, other protein molecules remaining in the solution after isolation and purification, as well as small molecular compounds) affect growth and are often abundantly incorporated into the crystals.
Several detailed studies were carried out with the proteins lysozyme and ferritin (or apoferritin). For both proteins the impurity species that remains in the solution even after laborious purification is a dimer of the native molecules, likely because of its chemical similarity to the native protein (Thomas et al., 1996, 1997, 1998).
With lysozyme, the dimer has an isometric shape and contains two covalently bound monomer chains. It was shown to severely affect the shape of the interface, reduce the growth rate by factors of up to 5, and completely inhibit growth at low supersaturations (Thomas et al., 1996, 1998; Vekilov, 1993; Vekilov et al., 1995; Vekilov and Rosenberger, 1996). It was found that the dimers are preferentially incorporated into the crystals and exhibit severely nonuniform distribution with a core of very high impurity incorporation in the central regions of a crystal (Stojanoff et al., 1997; Vekilov et al., 1996b). It was also found that impurities adsorbed on the growth interface cause severe growth unsteadiness and step bunching (Vekilov and Alexander, 2000; Vekilov et al., 1996a; Vekilov and Rosenberger, 1998a). The nonuniform impurity incorporation and the step bunching have been shown to cause high levels of lattice strain, structural nonuniformities and mosaicity (Caylor et al., 1999; Chernov, 1998, 1999; Dobrianov et al., 1998; Yau et al., 2001).
With ferritin, the "dimers" consist of two slightly misfolded native spheres held together by hydrophobic bonds (Petsev et al., 2000). As with lysozyme, they are preferentially incorporated into the crystals (Thomas et al., 2000). Each incorporated dimer causes stacks of trivacancies and single vacancies up to five molecular layers high (Yau et al., 2001, 2000b). The vacancies stress the crystal lattice, and the accumulated strain is resolved by the breaking of crystals larger than 100–200 µm into mosaic blocks of 50 µm in size (Yau et al., 2001).
The relatively large aggregates of evidence in Fig. 7 are similar to those for crystallization of ferritin and lysozyme in that they consist of the crystallizing protein. In further similarity, the impurities are incorporated into the crystals and severely affect growth. However, there are some significant differences. The HbC aggregates are large and contain multiple native or denatured molecules. While the lysozyme and ferritin dimers exist before crystallization and could, potentially, be removed with a high-sensitivity technique, the HbC aggregates form only after a high-concentration crystallization solution is prepared. Thus, they are a constant companion of crystallization and can only be avoided if a different set of crystallization conditions is employed. While it has been known (Ferre-D-Amare and Burley, 1994) that protein aggregation should be avoided for successful protein crystallization, it was thought that this is because aggregation provides an alternative pathway to lower the supersaturation of a concentrated protein solution. The above observations with HbC show that even when the aggregate concentration is too low and aggregation cannot be a real competition to crystal formation, aggregates can still interfere with the growth of perfect crystals.
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ACKNOWLEDGEMENTS |
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This work is part of the doctoral thesis of Angela R. Feeling-Taylor in partial fulfillment of the requirements for the doctoral degree in the Department of Anatomy and Structural Biology of the Albert Einstein College of Medicine.
This work was supported by the National Heart, Lung, and Blood Institute, National Institutes of Health, through grants HL58038 and HL3865, and National Institutes of Health Minority Predoctoral Fellowship 1F31 HL09564-01 to A.R.F.-T; the Office of Biological and Physical Research, National Aeronautics and Space Administration, through grants NAG8-1354 and NAG8-1854; and the American Heart Association, Heritage Affiliate grant-in-aid No. 9950989T.
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FOOTNOTES |
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Submitted on January 7, 2004; accepted for publication June 16, 2004.
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