Advances in Protein Chemistry Volume 41

ADVANCES IN PROTEIN CHEMISTRY Volume 41

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ADVANCES IN PROTEIN CHEMISTRY EDITED BY C. B. ANFINSEN

JOHN T. EDSALL

Department of Biology The Johns Hopkins University Baltimore, Maryland

Department of Biochemistry and Molecular Biology Harvard University Cambridge, Massachusetts

FREDERIC M. RICHARDS

DAVID S. EISENBERG

Department of Molecular Biophysics and Biochemistry Yale University New Haven, Connecticut

Department of Chemistry and Biochemistry University of California, Los Angeles Los Angeles, California

VOLUME 41

ACADEMIC PRESS, INC. Harcourt Brace Jovanovich, Publishers

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COPYRIGHT 0 1991 BY ACADEMIC PRESS, INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in Writing from the publisher.

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ISBN 0-12-034241-3 (ak.paper)

PRINTED IN THE UNITED STA'IES OF AMERICA 91

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CONTENTS

Physical Principles of Protein Crystallization PATRICIA C . WEBER

I . Introduction

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I1. Stagesof CrystalGrowth

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. I11. Driving Forces for Crystal Growth . IV. Nucleation . . . . . . V. Crystal Growth Mechanisms . .

VI . VII . VIII . IX . X. XI . XI1 .

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Competition between Nucleation and Growth Cessation of Growth and Crystal Disorder . Crystallization Methods . . . . . Protein Purity . . . . . . . Searching for Crystallization Conditions . . New Developments in Protein Crystallization . Summary Remarks . . . . . . References . . . . . . . .

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Protein Hydration and Function JOHN

A . RUPLEY AND GIORGIO CARERI

I . Introduction . . . . 11. Thermodynamics . . . I11 . Dynamics . . . . IV. Structure . . . . V. Computer Simulation . . VI . Picture of Protein Hydration VII . Hydration and Function . VIII . Conclusion . . . . Appendix: Percolation Theory References . . . .

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CONTENTS

Lysozyme and a-Lactalbumin: Structure, Function, and Interrelationships HUGHA. MCKENZIEAND FREDERICK H. WHITE,JR. I. Introduction .

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11. Early History. . . . . . . . . . 111. Some Aspects of the Occurrence, Isolation, and Characterization of Lysozyme and a-Lactalbumin . . . IV. Three-Dimensional Structure of Lysozyme . . . V. Three-Dimensional Structure of a-Lactalbumin . . VI. Comparative Binding of Metal Ions in Lysozyme and

174 176 181 192 206

a-Lactalbumin . . . . . . . . . VII. Amino Acid Composition and Sequence Homologies in Lysozymeanda-Lactalbumin . . . . . VIII. Galactosyltransferase and the Lactose Synthase System . . . . . . . . . . . IX. Some Additional Physical, Chemical, and Biological Comparisons between Lysozyme and a-Lactalbumin X. Evolutionary Origins of Lysozyme and a-Lactalbumin XI. Conclusions and the Future . . . . . . References . . . . . . . . . . Note Added in Proof . . . . . . . .

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AUTHOR INDEX

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224 250 259 276 293 299 315

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By PATRICIA C.WEBER

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Central Research & DevelopmentDepartment. E.1 du Pont de Nemoursand Co., Inc., Wilmington. Delaware 19880

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Stages of Crystal Growth . . . . . . . . . . . . . . . . . . . . . . . . 111. Driving Forces for Crystal Growth . . . . . . . . . . . . . . . . . . . . IV. Nucleation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A . Homogeneous Nucleation . . . . . . . . . . . . . . . . . . . . . . B . Heterogeneous Nucleation . . . . . . . . . . . . . . . . . . . . . . C. Nucleation Rate . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Experimental Determination of Nucleation Conditions . . . . . . . . V. Crystal Growth Mechanisms . . . . . . . . . . . . . . . . . . . . . . A . Transport-Controlled Growth . . . . . . . . . . . . . . . . . . . B . Growth Controlled by Surface Kinetics . . . . . . . . . . . . . . . C. Measurements of Crystal Growth Rates . . . . . . . . . . . . . . . D . Transport Phenomena in Protein Crystal Growth . . . . . . . . . . . E. Role of Molecular Preassociation in Nucleation and Crystal Growth . . . VI . Competition between Nucleation and Growth . . . . . . . . . . . . . . VII . Cessation of Growth and Crystal Disorder . . . . . . . . . . . . . . . . VIII . Crystallization Methods . . . . . . . . . . . . . . . . . . . . . . . . A . Batch Method . . . . . . . . . . . . . . . . . . . . . . . . . . . B . Dialysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Vapor Diffusion . . . . . . . . . . . . . . . . . . . . . . . . . . D. Temperature Shift . . . . . . . . . . . . . . . . . . . . . . . . . E . Achieving Different Conditions for Nucleation and Growth . . . . . . F. Free Interface Diffusion . . . . . . . . . . . . . . . . . . . . . . G. Seeding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX . Protein Purity . . . . . . . . . . . . . . . . . . . . . . . . . . . . X . Searching for Crystallization Conditions . . . . . . . . . . . . . . . . XI . New Developments in Protein Crystallization . . . . . . . . . . . . . . A . Crystallization in Microgravity . . . . . . . . . . . . . . . . . . . B . Automated Crystallization . . . . . . . . . . . . . . . . . . . . . XI1 . Summary Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I . INTRODUCTION

Protein crystals are three-dimensionally ordered arrays of biological macromolecules . Although the dimensions of these crystals that sparkle and polarize light are measured in only tenths of millimeters. their ability to diffract X-rays provides the experimental data needed to image 1 ADVANCES Ihi PROTEIN CHEMISTRY. Vol. 41

Copyright 0 1991 by Academic Press. Inc. All rights of reproduction in any form reserved .

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biological structures at atomic resolution. The detail that can be resolved by X-ray crystallography depends on the degree of molecular and lattice ordering in the crystal. In the absence of well-ordered crystals, X-ray studies at atomic resolution are impossible. Protein crystals are also used in neutron diffraction studies and a variety of optical and magnetic resonance spectroscopies. Here, the molecular order of the crystal enhances the directional resolving power of experimental methods that include Mossbauer, electron spin resonance, circular dichroism, and Raman spectroscopies. In an emerging technology, the ordered assembly of biological macromolecules is envisioned to allow the construction of new biomaterials (Furuno and Sasabe, 1985). Proteins are crystallized from aqueous solutions using methods that have been extensively studied for simpler molecules and salts (Rosenberger, 1986; Feigelson, 1988). Despite similar underlying physical principles, protein and small-molecule crystallizations differ in many respects. Unlike the crystallization of simpler molecules, in which solvent is effectively excluded from the crystal, substantial numbers of solvent molecules are immobilized and become ordered at protein lattice contacts, although otherwise protein crystals have large cavities containing essentially liquid water. An important feature of protein crystal growth experiments is the need to carry out crystallization trials with very small quantities of scarce and expensive materials. When experiments are carried out in such small volumes (typically, 5-100 pl), it becomes difficult to define and control solution properties. The situation becomes particularly complicated when vapor diffusion or other nonequilibrium approaches to crystal growth are used, as these produce different and changing conditions throughout the small volumes involved. This article reviews recent work on various aspects of the physical chemistry of protein crystal growth. Several books and reviews treat experimental and technical aspects of this area (e.g., Blundell and Johnson, 1976; McPherson, 1982, 1989; Michel, 1983; Sheshadri and Vankatappa, 1983; Matsuura, 1985; Wyckoff et al., 1985; Garavito et al., 1986; Ollis and White, 1989; see also the Proceedings of the First and Second International Conferences on Protein Crystal Growth [J. Cryst. Growth 76,535-718 (1986),J. Cryst. Growth 90, 1-374 (1988),and Carter, 19901. The objective here is to relate physical conceptions of how protein crystals grow in order to understand and improve existing crystallization methods. The ultimate practical goal is to allow the easy crystallization of targeted proteins in order to realize the potential utility of structural knowledge in protein engineering and drug design.

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N

Crystal

Associated Chains

FIG. 1. Reversible molecular association reactions involved in the assembly of crystals. Monomers initially combine into small aggregates (here, called chains). The association of monomers into chains leads to the formation of prenuclear aggregates that continue to grow by further addition of monomers or chains. The partition of molecules into monomers, chains, and prenuclear aggregates is called a quasiequilibrium state (Kam et al., 1978). When sufficient molecules associate in three dimensions, a thermodynamically stable critical nucleus is formed. The addition of monomers and/or chains to critical nuclei eventually leads to the formation of macroscopic crystals.

11. STAGES OF CRYSTAL GROWTH

Crystallization is a complex multiequilibrium process (Fig. 1). The three stages of crystallization common to all molecules are nucleation, crystal growth, and cessation of growth. During nucleation enough molecules associate in three dimensions to form a thermodynamically stable aggregate. These nuclei provide surfaces suitable for crystal growth. Crystal growth ceases when the solution is sufficiently depleted of protein molecules, deformation-induced strain destabilizes the lattice, or the growing crystal faces become poisoned by impurities. 111. DRIVING FORCES FOR CRYSTAL GROWTH

Crystals form in supersaturated solutions in which the solute concentration exceeds its solution solubility. Supersaturation is usually expressed as either of the ratios cIc, or (c - cs)Ic,, where c is the concentration of solute before crystallization and c, is the solute equilibrium saturation concentration. Supersaturated solutions are thermodynamically metastable. Equilibrium can be restored by reducing the solute concentration through precipitation or formation of nuclei and subsequent crystal growth. The supersaturation requirements for nucleation and

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PATRICIA C. WEBER

C

0 .c

s

c

C

0 ( I) S 0

0

.-C(I) c 9

a

.1 Curve + Solubility-Decreasing

Parameter

FIG. 2. A hypothetical protein solubility graph showing the changes in supersaturation for commonly used protein crystallization methods. Protein concentration is plotted as a function of a parameter that decreases protein solubility. The solubility curve divides the solubility graph into supersaturated and unsaturated regions. Supersaturated solutions support crystal growth, with increased rates observed at higher supersaturation levels (smaller abscissa values). At supersaturation levels greater than the supersolubility curve, homogeneous nucleation occurs (after Feigelson, 1988). Point A shows the supersaturation level of a batch crystallization experiment in which the protein solution is mixed with precipitating agents to achieve supersaturation and then left unchanged. The change in protein supersaturation during typical vapor diffusion experiments is shown by the line from B to C. Solutions are unsaturated on setup (B). During equilibration the solution enters the supersaturated region (C). Nuclei form when the supersaturation exceeds the supersolubility curve. If the supersaturation is then lowered by moving from C to D, only larger stable nuclei remain to support crystal growth. In a free interface diffusion experiment, when the protein and precipitant solutions are first layered, molecules at the proteinprecipitant interface are sufficiently supersaturated to spontaneously nucleate (E). The remaining protein solution is unsaturated (F). On equilibration, the entire protein solution is supersaturated (G).

growth are different (Fig. 2). For a given solute, spontaneous nucleation occurs at high supersaturation, whereas lower supersaturation will support growth of a seed crystal, but not spontaneous nucleation. At concentrations below saturation, crystals dissolve. Protein crystal growth involves the incorporation of a complex unit into an existing lattice. The growth unit usually includes the covalent polypeptide chain, water molecules that are integral components of the folded protein structure, and additional water molecules and solvent ions that may become immobilized at crystal lattice contacts. Direct inter-

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

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actions between protein molecules are relatively tenuous in most protein crystals that have been examined in detail (Frey et al., 1988; Salemme et al., 1988). Typically, water molecules that become immobilized during crystal formation serve to fill irregular gaps that occur between molecules at lattice contacts. Occasionally, intermolecular salt linkages (Baker, 1988; Dreusicke et al., 1988) or counterions (Sheriff et al., 1987) form electrostatic interactions at crystal contacts to stabilize the lattice structure. Crystals are entropically destabilized, owing to both the loss of rigidbody molecular translational and rotational degrees of freedom and the immobilization of surface loops that may be flexible in solution, but become ordered at lattice contacts (Finzel and Salemme, 1985; Sheriff et al., 1985; Salemme et al., 1988). Although some immobilization may be a natural consequence of packing objects that tend generally to have loops on their surfaces, loop flexibility may more easily accommodate minor structural changes that facilitate incorporation of the protein into a crystalline lattice (Salemme et al., 1988). Although losses of molecular entropy make unfavorable contributions to the stabilization free energies of lattice formation, some of this stabilization can be recovered due to the appearance of lattice vibrational modes, evidence for which is seen from some protein crystal studies (Finzel and Salemme, 1986; Caspar et al., 1988). Proteins are generally induced to crystallize by adding agents that either alter their surface charges, or perturb the interactions between the protein and bulk solvent water to promote associations that lead to crystallization. While many proteins crystallize near their isoelectric points in low ionic strength solutions (Blundell and Johnson, 1976), it is more common to use organic molecules, polymers, or salts at high concentrations to promote crystal growth. Most “precipitants” change the chemical potential of the protein in solution and act by affecting the partition of water between the protein and the precipitant (Timasheff and Arakawa, 1988). The protein usually has a higher affinity for water than it has for the precipitant. The preferential interaction with water creates a precipitant-poor layer near the protein surface (Fig. 3). Formation of the exclusion layer is thermodynamically unfavorable. Protein association is favored because it decreases the area of the precipitant-poor layer near the protein surface (Fig. 3). Conversely, the additives may also stabilize protein structure by concentrating water near the protein surface and possibly favoring more compact structural organizations (Arakawa and Timasheff, 1984). Random-chain polymers (e.g., polyethylene glycol) are also frequently used to promote crystallization. These polymers act to preferentially hydrate the protein through excluded volume effects,

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Additive-depleted Layer

FIG. 3. Preferential protein hydration in the presence of precipitating agents used in crystallization experiments. When high concentrations of salts are used as precipitants, a precipitant-poor layer forms near the protein (P) surface due to a higher affinity of the protein for water than for the precipitant. Other precipitants (e.g., polyethylene glycol polymers) induce formation of a similar precipitant-depleted region near the protein by solvent exclusion effects. In either case formation of the precipitant-depleted layer is energetically unfavorable. Consequently, the overall effect of precipitants is to promote molecular associations that decrease the total protein surface area exposed to solvent. After Timasheff and Arakawa (1988).

whereby the extended chain polymer and its entrained water are excluded from the area near the protein (Arakawa and Timasheff, 1985). Similar to the situation with salts, this system becomes energetically more favorable when the protein molecules associate to minimize unfavorable surface tension effects. IV. NUCLEATION A. Homogeneous Nucleation

The smallest stable unit of a crystal is the nucleus. Nuclei are formed by either homogeneous or heterogeneous nucleation. Homogeneous nucleation is the spontaneous formation of solute nuclei in a supersaturated solution. In the absence of external changes, the force for spontaneous nucleation arises from fluctuations in solution. The energy required to form stable nuclei from monomeric species in solution is the sum of opposing free-energy terms. With increasing incorporation of molecules, the nucleus becomes more stable as favorable intermolecular contacts form in the three-dimensional lattice. However, formation of the nuclear surface produces an energetically unfavorable surface tension contribution. The incremental increase in surface tension on the addition of molecules to the nucleus becomes smaller as nuclei become

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

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larger. Consequently, beyond a critical size, the energetically favorable volume term becomes dominant and nuclei are stable. The activation energy of nucleation decreases at higher supersaturation and increased temperatures. The critical nuclear size (i.e., the number of molecules needed to form a stable nucleus) also decreases with increasing supersaturation (Kam et al., 1978; Boistelle and Astier, 1988).

B . Heterogeneous Nucleation Heterogeneous nucleation is the formation of solute nuclei on foreign substrates such as dust particles or surface irregularities in the container. The activation energy for heterogeneous nucleation is less than that for homogeneous nucleation, due to an attraction between the solute and the nucleant, so that heterogeneous nucleation occurs at lower supersaturation. Frequently, crystals grow on foreign nucleation sites and never appear in the bulk solution. Although the nucleation activation energy is lower for heterogeneous nucleation, the critical dimensions are similar for nuclei formed by heterogeneous and homogeneous nucleation mechanisms (Boistelle and Astier, 1988). Although heterogeneous nucleation of protein crystals frequently occurs accidentally, systematic studies of nucleation on mineral substrates demonstrate successful protein crystal growth (McPherson and Shlichta, 1988) (Fig. 4A). Several different minerals were tested with each of several proteins. Usually, a given protein crystallized on only a subset of minerals, indicating that heterogeneous nucleation involved some fairly specific interaction between the protein molecules and the mineral surface. Nucleation and crystal growth occurred more rapidly and at lower supersaturation than in the absence of the mineral nucleant. Interestingly, epitaxial growth was observed for lysozyme on the mineral apophyllite (McPherson and Shlichta, 1988). In this case a surface of the apophyllite crystal presents a two-dimensional lattice repeat that is a nearly exact fraction of the lysozyme cell dimensions. C . Nucleation Rate

Crystal nucleation rates, expressed as the number of nuclei formed per unit volume per unit time, increase with protein solubility. Higher solubility leads to increased molecular encounters in solution and reduced levels of supersaturation required for spontaneous nucleation. Nucleation rates typically show a high-power dependence on protein supersaturation, and so empirically increase rapidly above a critical value

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of protein supersaturation (Boistelle and Astier, 1988). At supersaturation levels below the critical value, the slow nucleation rates lead to long times between attaining supersaturation and achieving nucleation. The slight difference between supersaturation levels where nucleation is very slow and very fast makes the nucleation rate difficult to control experimentally. The exponential dependence of nucleation rate on supersaturation has been studied in detail for sickle cell hemoglobin (hemoglobin S) (Hofrichter et al., 1974, 1976) and tetragonal lysozyme (Ataka and Tanaka, 1986). In both cases the elapsed time prior to crystal appearance depended on a high power of supersaturation. For hemoglobin s, nucleation rates were also significantly faster at higher temperatures

FIG. 4. Crystal photographs. (A) The heterogeneous nucleation of Streftomyces avzdzniz streptavidin crystals on the mineral biotite. (B) Streptavidin crystals are shown growing at the surface of a hanging drop. (C and D) Pseudomonm indigofera isocitrate lyase crystals grown (C) on earth and (D) in microgravity aboard the STS-26 space shuttle. (E-H) Crystals grown using an automated pipetting device are shown. Crystals of recombinant human interleukin lp. [(E) Gilliland et al. (1987); D. B. Carter et al. (1988)l and apostreptavidin [(F) Pahler et al. (1987)] were reproduced from conditions reported in the literature. Crystallization conditions for (G) E. coli ketol-acid reductoisomerase and (H) a Fab fragment of a monoclonal antibody to angiotensin were found using successiveautomated grid searches (Cox and Weber, 1988). Bar (A-H): 0.1 mm.

FIG.4B and C.

FIG.4D and E. See legend on p. 8 10

FIG.4F and G. See legend on p. 8. 11

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PATRICIA C. WEBER

FIG.4H. See legend on p. 8.

(Hofrichter et al., 1976), again reflecting the dependence of the nucleation rate on the frequency of molecular encounters. D . Experimental Determination of Nucleation Conditions

Experimental methods for the early detection of nuclei formation using solution light-scattering measurements were described by Kam et al. (1978). A principal difficulty is to distinguish between the formation of three-dimensional nuclei and amorphous aggregates at early stages of protein association. Kam et al. (1978) developed a two-parameter model to discriminate between these processes, based on the idea that the number and distribution of intermolecular contacts formed by nuclei and precipitates differ. Nuclei are compact, each molecule making several three-dimensional intermolecular contacts, while precipitates form more extended chain networks that are larger and less dense. This variation in size and density gives rise to different signals in dynamic lightscattering measurements as a function of protein concentration and allows discrimination between the two aggregation states prior to macroscopic crystal formation (Kam et al., 1978; Feher and Kam, 1985). This

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

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approach has been used to study nuclei formation of lysozyme (Kam et al., 1978; Baldwin et al., 1986) and phosphoglucomutase (C. W. Carter et al., 1988).

V. CRYSTAL GROWTH MECHANISMS A . Transport-Controlled Growth Crystal growth rates depend potentially on both the transport rates of solution molecules to the crystal surfaces and their rate of incorporation after they have arrived. Models of crystal growth (Fiddis et al., 1979; Davey, 1986; Boistelle and Astier, 1988) have been developed that distinguish between transport and surface-ordering events as factors that control growth rates. In the transport-limited growth model, growth rates reflect the frequency with which molecules reach the crystal surface. Although many growth experiments show evidence for a depletion region around growing crystals (Kam et al., 1978; Pusey et al., 1988), as described in Section V.D, most studies suggest that surface effects are rate limiting in protein crystal growth.

B . Growth Controlled by Su7face Kinetics Experimental studies of lysozyme (Fiddis et al., 1979; Pusey et al., 1986), insulin (Schlichtkrull, 1957; Fiddis et al., 1979), and canavalin (DeMattei and Feigelson, 1989) suggest that events occurring at the lattice surface are rate limiting in protein crystal growth. Important effects occurring at the lattice surface include the formation of favorable growth sites (usually some form of irregularity) and molecular attachment to these sites. For rough crystal surfaces, where many growth sites exist or the energy to create them is low, growth proceeds at relatively low values of supersaturation. Growth on smooth surfaces, where the energy to create a growth site is high and may necessitate the formation of surface nuclei, requires relatively higher supersaturation levels. In more detailed terms the growth models include 1. A rough surface model in which molecular incorporation is favored at many vacant sites in the nascent lattice layer. In this case the growth rate is G = k,(c -

cS)

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PATRICIA C. WEBER

where k, depends on the surface binding energy and the mass transfer rate to the crystal surface, c is the protein solution concentration, and cs is the protein solubility. 2. A screw dislocation model that predicts preferred growth along a defined dislocation. Here,

G

=

k,c(ln c/cS)*

where k, similarly depends on characteristics of the growing crystal face. 3. A surface nucleation model in which attachment sites exist as molecular clusters that, like nuclei, must reach a critical size to be stable and support subsequent crystal growth. Here, G = k3c1’3(lnC / C ~ ) ~exp[ ’ ~ - ~ y ‘ / 3 ( k ~ T )In ’ c/cs]

where k, is a function of the crystal face, y is the excess free energy of a molecule with unsatisfied lattice interactions at the edge of a growth site, k, is Boltzmann’s constant, and T is the temperature. Because these models predict various dependencies of the crystal growth rate on solution supersaturation, growth mechanisms can be distinguished experimentally (Schlichtkrull, 1957; Fiddis et al., 1979; Durbin and Feher, 1986; Pusey et al., 1986; DeMattei and Feigelson, 1989). Most of these studies suggest that surface nucleation and screw dislocation models most accurately describe protein crystal growth kinetics, although a detailed study of hen egg white lysozyme (Durbin and Feher, 1986) shows different mechanisms at low and high protein supersaturation levels. At low supersaturation different growth rates were observed on equivalent faces of tetragonal lysozyme crystals. This result suggested that preferred growth occurs at a very small number of local surface defects, presumably introduced at random to account for unequal growth rates of equivalent crystal faces. At higher supersaturation equivalent faces of lysozyme crystals grow to similar size. In this case the growth rate dependence on supersaturation follows the two-dimensional nucleation model. C . Measuremnts of Crystal Growth Rates Growth rates on the order of 10-8 cm/sec have been measured for several protein crystals (Fiddis et al., 1979; Pusey et al., 1986; DeMattei and Feigelson, 1989). In general, crystals grow faster at increasing levels of supersaturation, and except for small crystals (i.e., < 10 pm), growth rates appear to be independent of size (Schlichtkrull, 1957). In a comparison of protein and small-molecule crystal growth rates from solution,

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

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Feigelson (1988) found that their surface linear growth rates are comparable. When the number of molecules added per unit time is considered, protein crystals do grow more slowly. However, because protein molecules are much larger, fewer are needed to achieve the linear growth rates observed for small molecules.

D. Transport Phenomena in Protean Crystal Growth Sustained crystal growth requires that solute molecules continually reach the crystal surface. Transport of molecules can occur by both diffusion and convection. Because the solution near the growing crystal surface is depleted of solute as the crystal grows (Fig. 5 ) , gravity can act on the density difference between the solute-depleted layer and the bulk solvent to produce convection currents. Although many studies of crystal growth describe the transport phenomenon responsible for mass transfer to growing crystals as “diffusion,” theoretical arguments suggest that buoyant or, in small volumes, surface tension-driven convection actually dominates simple Fick’s law diffusion in determining the

I Distance

-

FIG. 5. Protein, protein contaminant, and salt concentrations near the surface of a growing crystal. As molecules add to the crystal, the solution near the crystal is depleted of protein. The exclusion of protein contaminants and salts increases their effective concentration near the crystal surface. The relative shapes of the concentration curves depend on the molecular diffusivity, with more rapidly moving molecules such as salts having wider concentration gradients. The concentration profiles shown are expected to occur in the absence of convection. Convection currents caused by the differences in solution density greatly diminish the extent of the Concentration gradients.

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rates of mass transfer to growing crystals (Rosenberger, 1986). Indeed, buoyancy-induced convection currents around growing protein crystals have been directly observed using schlieren optics that allow visualization of variations in the solution-refractive index (Pusey et al., 1988). Pusey et al. (1986) suggest from their data on lysozyme that convection due to density gradients near the growing crystal is sufficient to prevent diffusion-limited crystal growth. Flow of solute also influences protein crystal morphology. Preliminary experiments show that relative growth rates of human serum albumin crystal faces depend on crystal orientation in a flowing solution (Broom et al., 1988). Based on the observed changes in relative sizes of crystal faces, these authors speculate that unfavorable habits such as needles could be improved by oriented seeding.

E . Role of Molecular Preassociation in Nucleation and Crystal Growth Models for protein crystal formation follow those for small molecules and assume that crystal nuclei form and grow by the association or addition of solute monomers. Nevertheless, several lines of evidence suggest that aggregates may participate in some aspects of protein crystal nucleation and growth. It is a common experience that the onset of nucleation and crystal growth are delayed for long periods of time after suitable supersaturating conditions exist. Kam et al. (1978) suggest that this preequilibrium state is characterized by the formation of various molecular aggregates prior to the eventual formation of stable nuclei. In fact, solution studies by Banerjee et al. (1975) show that lysozyme self-associates into indefinite head-to-tail polymers under conditions similar to those used for crystallization experiments. Analysis of molecular interactions in several lysozyme crystal forms (Salemme et al., 1988) showed that the polymorphs could all be assembled from a common subset of linear molecular chains (Fig. 6). Since the crystals and polymers form under similar conditions (Banerjee et al., 1975), it is possible that chains observed in solution represent preaggregates that associate to form crystal nuclei. Such “sequential” mechanisms might provide easy formation routes for protein crystal nuclei where the molecules are only tenuously connected in the threedimensional lattice. Whether crystals can grow by the addition of molecular aggregates (as opposed to single molecules) to the crystal faces is less clear. T h e observation that the predominant growth mechanisms for many crystals involve two-dimensional surface nucleation (Schlichtkrull, 1957; Fiddis et al., 1979; Durbin and Feher, 1986; Pusey et al., 1986) could be the result of molecular chain association to an otherwise smooth and com-

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b FIG. 6. Stereoscopic views of common molecular chains in three crystal forms of hen egg white lysozyme. (a) Views of monoclinic (A), tetragonal (B), and triclinic (C) cells, illustrating the recurrence of the chain corresponding to the triclinic c axis. (b) The triclinic c-axis chain (A) aligned with one subunit of the dimer in the asymmetric unit of the monoclinic cell (B). (C) The triclinic a-axis array oriented with the other subunit of the monoclinic asymmetric unit. From Salemme et al. (1988).

pleted lattice plane. In this case growth rates could depend on the polymeric species distribution in complicated ways. For lysozyme the heat of formation for the self-associated chains (Banerjee et al., 1975) is -6.4 kcal/mol, a value comparable to measured heats of crystallization for tetragonal lysozyme [ - 17.2 kcal/mol (Ataka and Asai, 1988), and - 25.1 kcal/mol (Takizawa and Hayashi, 1976)], although these apparently vary somewhat, depending on solution conditions. VI. COMPETITION BETWEEN NUCLEATION AND GROWTH Rates of crystal nucleation and growth generally have different dependencies on protein supersaturation, and additionally vary substantially for different protein-precipitant systems. This can lead to a variety of unexpected behaviors in crystallization experiments. T h e nucleation

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rate is typically an exponential function of supersaturation, ranging from values of 3-4 for lysozyme (Ataka and Tanaka, 1986) to 35 for hemoglobin S (Hofrichter et al., 1976). Growth rates, in contrast, can have a variety of functional forms, although supersaturation levels that support nucleation also generally promote rapid growth. T h e difference in the dependence of nucleation and growth rates on supersaturation has important implications for experiments in which the desired result is the formation of a few large crystals. For example, in systems in which the nucleation rate depends on a relatively low power of supersaturation, narrow ranges of supersaturation exist that favor growth over nucleation (Ataka and Tanaka, 1986). For these systems nucleation is likely to dominate at low and high supersaturations, while crystal growth is favored at intermediate supersaturation. This model is supported by data from several systematic studies of crystal size and number as a function of supersaturation. For rabbit muscle aldolase (Heidner, 1978) and hen egg white lysozyme (Ataka and Tanaka, 1986), more crystals formed at high and low supersaturation and fewer were observed at intermediate supersaturation levels. In studies of crystal size as a function of protein concentration (Heidner, 1978; Ataka and Tanaka, 1986; Betts et d.,1989), final crystal size depended critically on the supersaturation ratio, the ratio of initial protein concentration to protein solubility. The linear dimensions of lysozyme crystals, for example, could be increased from 0.6 mm to 1.0 mm, using solutions in which the supersaturation ratio varied from 2.5 to 4 (Ataka and Tanaka, 1986) (Fig. 7). Betts et al. (1989) also obtained large crystals (1.0 X 0.6 1.o

a, N

i7, 0.5

n "

1

2

4

3

5

6

c/c s FIG. 7. The relationship between final size of hen egg white lysozyme crystals and the degree of supersaturation. The largest crystals grew when the ratio of initial lysozyme concentration (c) to its solubility (c,) ranged between 2.5 and 4.0. From Ataka and Tanaka (1 986).

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

19

x 0.4 mm) of cytidine deaminase over a narrow range of protein concentrations. At protein concentrations above and below the optimum, only precipitate formed or else the solution remained clear. OF GROWTH AND CRYSTAL DISORDER VII. CESSATION

Perhaps the most practical, yet mysterious, aspect of protein crystallization concerns the causes of growth cessation. It has frequently been observed that some crystals do not grow beyond a certain size, even in the presence of excess protein. There are several possible reasons for this effect. Probably the most common, in view of the complexity and chemical sensitivity of protein molecules, is the gradual poisoning of the crystal surfaces by defective molecules that themselves attach to the crystal lattice, but do not support subsequent growth. If the defective molecules bind more weakly than native molecules, then they tend to be excluded from the lattice until the native molecules are nearly depleted from the solution (Fig. 5). At this stage, owing to the higher solution concentration of the defective molecules, their addition to the lattice dominates and the surfaces become poisoned toward further growth. For example, crystallization of hemoglobin C is inhibited by the addition of hemoglobins A and F (Hirsch et al., 1988). Although it is apparent that progressive poisoning of crystals is possible (and probably underlies the strain-defect cessation model described below), crystal growth can occasionally be reinitiated by changes in the surrounding protein solution (Young et al., 1988), suggesting the localization of the poisoning molecules at the crystal surface. Similarly, macroseeding experiments (Section VII1,G) are usually initiated with an etching step that presumably removes defective surface molecules that would otherwise poison the crystal surfaces toward further growth. An alternative mechanism that can lead to growth cessation is the introduction of crystal strain or cumulative defects into the lattice as the crystal grows. The concept was physically realized by Kam et al. (1978), who halved a lysozyme crystal and found that each half then grew to the size of the original. It was suggested that this reflected the continuous accretion of defects while the crystal grew, so that finally the addition of new molecules to the (defective) surface lattice became unfavorable. Although the incorporated defects could be either molecular or structural (resulting, say, from too rapid growth), the observation that the crystal halves regrew to the original size suggests that the defects were structural, since they were propagated into the newly reformed lattices. Structural defects might be expected to be more common in rapidly grown crystals, as suggested by many experiments showing that growth rates can affect terminal growth size. For example, under otherwise simi-

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lar conditions, larger crystals of hen egg white lysozyme were grown in hanging-drop experiments when supersaturation was achieved slowly at controlled rates (Gernert et al., 1988). Although the effects of incorporating defective molecules into the growing crystal lattice have been introduced in the context of growth cessation, it is clear that situations can exist in which the crystal can continue to grow macroscopically, even though it incorporates defects that destroy the long-range molecular order in the lattice. The disappointing result is a crystal that typically looks good, may be strongly birefringent, but does not usefully diffract X-rays. The variety of diffraction effects observed-ranging from no diffraction, through resolution-limited or anisotropic diffraction patterns, to disordered patterns in which still exposures look like precession photographs-illustrates the many types and spatial scales of lattice disorder that can occur (Harburn et al., 1975).

METHODS VIII. CRYSTALLIZATION The commonly used protein crystallization methods achieve and maintain supersaturation in several ways (Fig. 2). Severalarticles and books that describe methods used to grow protein crystals are referred to in Section I. The objective in this section is to briefly review methods as they relate to the phenomena described above. Examples of proteins crystallized by each method are given. More complete listings of crystallized proteins are compiled in McPherson, 1982 and Gilliland and Bickham, 1990. A . Batch Method

The simplest technique used to grow protein crystals is the batch method in which the protein is mixed with salts or other precipitants to achieve supersaturation (Fig. 2), and the vessel is sealed and set aside until crystals appear. Frequently, the supersaturation point required to induce nucleation is empirically determined by observing the onset of transient turbidity as powdered salt is progressively added to the solution. Crystals of hen egg white lysozyme used for most systematic studies of protein crystallization are grown by batch methods (Blundell and Johnson, 1976). Mouse pancreatic ribonuclease (Perry and Palmer, 1988)and the biotin operon repressor (Brennan et al., 1989) represent recent examples of use of the batch method. B . Dialysis

In the dialysis method protein solution is retained by a dialysis membrane which maintains the solution at a constant concentration while al-

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

21

lowing equilibration with a surrounding solution. Although the method can be used for any of the usual solvent perturbation approaches that involve added salts or small organic molecules to reduce protein solubility, the method is uniquely suited to the formation of crystals that are induced to crystallize at low ionic strengths. Occasionally, nucleation may be transiently induced at the dialysis solution boundary, where the membrane serves as a site for heterogeneous nucleation. Examples of the use of this method include the crystallizations of hexokinase (Steitz, 1971) and anthranilate phosphoribosyltransferase (Edwards et al., 1988).

C . Vapor Diffusion Vapor diffusion methods are among those most commonly used for protein crystallization because they readily lend themselves to the use of 5- to 5O-pl solution volumes. Typically, a hanging or sitting drop, containing a solution of protein plus precipitant at subsaturating concentrations of protein, is equilibrated against a larger reservoir of solution containing precipitant or another dehydrating agent. After sealing in a closed vessel, the solutions equilibrate to achieve supersaturating concentrations of protein and thereby induce crystallization in the drop. Theoretical models of hanging-drop experiments suggest that vapor equilibration at the droplet surface is sufficiently rapid to produce transient concentration gradients in the droplet (see, e.g., Yonath et al., 1982) (Fig. 4B) that might induce homogeneous nucleation (Fehribach and Rosenberger, 1989). Experimental measurements indicate final water equilibration times of 36-80 hr that depend on drop size and geometry and on reservoir precipitant concentrations (Fowlis et al., 1988; Mikol et al., 1989). When ammonium sulfate solutions are used in the reservoir, protein solution pH also rapidly equilibrates, owing to the low vapor pressure of ammonia (Mikol et al., 1989). Many additional variations are possible, particularly in systems including organic solvents in which diffusion both to and from the drop and the reservoir becomes a possibility. Controlled pH changes can be made by vapor equilibration of volatile organic acids or bases (McPherson and Spencer, 1975). D. Temperature Shift Proteins that are near their supersaturation points in concentrated salt solutions can frequently be induced to crystallizeby changing the temperature. This phenomenon, which has been used for the fractional purification of proteins (Jacoby, 1968), has found only scattered application in protein crystaHization. Nevertheless, given the relative ease of precise temperature regulation, methods based on temperature alteration de-

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PATRICIA C. WEBER

serve more thorough investigation (Rosenberger and Meehan, 1988). Clear applications include the introduction of temperature shifts to achieve transient nucleating conditions or to reduce the number of nuclei formed on initial solution supersaturation. Examples of proteins crystallized using temperature shift methodology include bovine neurophysin I1 (Rose et al., 1988) and elongation factor Tu (Lippmann et al., 1988). E . Achieving Different Conditionsfor Nucleation and Growth

The objective of most protein crystallization experiments is to obtain a few large crystals. As outlined in Sections IV,C and VI, two of the major obstacles to controlled protein crystal growth are the extreme sensitivity of nucleation rate to supersaturation conditions and the necessity for higher supersaturations to promote nucleation than are needed for growth (Fig. 2). An inherent shortcoming of many crystallization methods is that they depend on similar conditions both to promote nucleation and to support growth. A frequent result is either no crystals or the formation of many small crystals. However, alternative approaches have been developed that attempt to individually optimize nucleation and growth conditions. F. Free Inte$ace Diffusion Crystallization using free interface diffusion represents an attempt to achieve transient highly supersaturating conditions required for nucleation, followed by relaxation to conditions of lower supersaturation required for growth, within a single experimental setup (Salemme, 1972). In this method, a protein solution is layered over a precipitant solution. Initially, molecules at the liquid-liquid interface achieve high supersaturation, while the remainder maintain bulk conditions of the protein layer. The high supersaturating conditions at the interface promote nuclei formation. As the liquids diffuse, the high protein supersaturation initially achieved at the interface decreases. At equilibrium, when the precipitant and protein solutions are mixed completely, the entire protein solution is supersaturated (Fig. 2). Ideally, smaller nuclei dissolve at the lower levels of protein supersaturation and only the larger nuclei continue to grow. Cytochrome c ' (Weber and Salemme, 1977), phospholipase A2(Dijkstra et al., 1978),and adenylate kinase (Althoff et al., 1988) are among the proteins that have been crystallized by free interface diffusion.

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

23

G . Seeding

Seeding is a method that physically separates the processes of crystal nucleation and growth, so that conditions for crystal growth can be independently optimized. Seeding from crushed crystals can be done by using a fine glass rod to transfer nuclei from a stabilizing solution to a growth-promoting solution. An alternate microseeding method involves first passing a hair through a solution containing crystals or particles too small to be definitively identified as crystals, and then streaking the hair through the protein solution to be seeded (Stura and Wilson, 1990; Leung et al., 1989). Systematic methods of microseeding have been investigated (Fitzgerald and Madsen, 1986), as it is frequently difficult to control the number of seeds transferred via the glass rod or by streaking. Individual crystals having dimensions as small as 0.01 mm can be grown to larger sizes by macroseeding (Thaller et al., 1981). In this procedure a single crystal is repeatedly transferred to a fresh protein solution after crystal growth ceases. Before transfer to the new protein solution, the crystal is washed and its surface is etched by partial dissolution in a solution of low supersaturation. The need for etching suggests that crystal terminal size is caused by poisoning the crystal surface with impurities.

IX. PROTEIN PURITY It has long been recognized that protein purity plays a critical role in crystallization. Many investigators assert that if a protein fails to crystallize, or crystallization is irreproducible, the protein sample is simply lacking in sufficient purity (Anderson et al., 1988; Giege et al., 1988). As outlined in Section VII, impurities structurally similar to the solute are most likely to poison crystal growth or otherwise disrupt crystalline order. Both the availability of newer separation methods and the necessity to improve crystal quality in order to obtain key structural information have motivated detailed studies of how protein heterogeneity affects crystal growth. Protein contaminants can occur as natural isoforms or can arise during purification. Adventitious proteolysis and cysteine oxidation are probably the most common sources of microheterogeneity that occur during isolation (Lorber et al., 1987). This has frequently motivated the inclusion of protease inhibitors and/or reducing agents in crystallization solutions, as well as during purification. In many cases modifications that produce molecular heterogeneity are reflected in enzyme activity. For

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PATRICIA C. WEBER

example, it was noted that only the most active preparations of ribosome subunits would form useful crystals (Yonath et al., 1982). In many cases proteins which are otherwise intractable can be crystallized in fragments or as truncated forms. However, the chemical or enzymatic methods to cleave the molecules are a frequent source of product heterogeneity. For example, Fab fragments are liberated from intact immunoglobin molecules by endoproteolytic cleavage. The protease treatment often produces molecules having variability in the location of the cleavage site and extraneous nicks elsewhere in the molecule. The resultant Fabs are similar in structure, but differ in their isoelectric points. Purification of a single Fab isozyme by ion-exchange chromatography (Cygler et al., 1987; Boodhoo et al., 1988; Orbell et al., 1988), isoelectric focusing (Bott et al., 1982), or chromatofocusing (Prasad et al., 1988), followed by crystallization of the isoelectrically pure Fab fragment, has been shown to dramatically improve crystal quality. Similar results have also been reported as a result of separating isozymes of other biological macromolecules (Bott et al., 1982; Spangler and Westbrook, 1989). The complex consequences of cysteine oxidation were thoroughly detailed in a study of 4-hydroxybenzoate 3-monooxygenase prompted by an initial failure to reproduce the original crystal form (Van Der Laan et al., 1989). This work showed that enzyme crystallization was sensitive to the oxidation state of a single cysteine residue. Cysteine- 116, located at the molecular surface, is also situated at a lattice contact near Cys- 116 from an adjacent molecule. Consequently, crystallization of an enzyme having the cysteine oxidized to sulfonic acid is inhibited by the requirement to juxtapose two large negatively charged groups at a lattice contact occupied by reduced cysteines in the native crystals. It was additionally noted that only slight molecular movements could result in intermolecular disulfide formation. However, 4-hydroxybenzoate 3monooxygenase preparations contaminated with covalently linked aggregates form poorly ordered crystals, indicating the disruptive effect of this small contact modification on crystal lattice ordering. Although protein microheterogeneity usually disrupts crystal formation, it can occasionally promote crystallization. For example, crystals of Escherichia coli single-stranded binding protein contain a 1 : 1 mixture of intact and proteolyzed protein (Ollis et al., 1983). While crystallization experiments were initially conducted with intact protein, crystals grew only when enough molecules to form the mixed crystals had been cleaved by contaminating proteases. Recombinant DNA technology has had an enormous impact on crys-

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

25

tallography because it has made naturally scarce proteins plentiful. Moreover, it has largely eliminated proteolysis as a source of microheterogeneity in protein fragments, since these can now be produced genetically. A recent example of crystals grown from a genetically engineered fragment is the Klenow fragment of DNA polymerase (Brick et al., 1983). Nevertheless, biotechnological production methods can also introduce contaminants, particularly when the host organism expresses a protein similar to the foreign gene being overexpressed. In the expression of Bacillus stearothermophilus tryptophanyl-tRNA synthetase (trpS; tryptophan-tRNA ligase) in E. coli, low levels of E. coli trpS enzyme copurified with the cloned trpS and inhibited crystal growth. Only when the E. coli gene was deleted did the overexpressed B . stearothermophilus enzyme crystallize successfully (Carter, 1988). Many of the naturally scarce proteins that are typically overexpressed in E. coli functionally bind nucleic acids. Microheterogeneity often arises from nonspecific binding of host cell DNA or RNA, necessitating special isolation procedures to produce uncontaminated protein (Ruff et al., 1988).

X. SEARCHING FOR CRYSTALLIZATION CONDITIONS Although an extensive and expanding database of protein crystallization conditions reveals some trends in the uses of techniques and precipitants (Gilliland and Davies, 1984; Gilliland, 1988; Gilliland and Bickham, 1990), it is not yet possible to predict the conditions required to crystallize a protein from its other physical properties. Lacking any predictive scheme, the crystallization of a new protein is usually attacked using a more or less ad hoc approach based on the previous experiences of an investigator. Most often, crystallization attempts must be made with limited amounts of material, leaving the experimenter with the problem of searching a potentially large parameter space with a limited number of experiments. Despite, or perhaps owing to, these limitations, there have been two systematic approaches to searching for protein crystallization conditions. The first method to be described was an incomplete factorial approach (Carter and Carter, 1979). This is basically a method that, given a matrix of compositional components and their concentrations, defines how to sample the variables with a minimum number of experiments. Using statistical methods to analyze the results, it is possible to identify variables that are correlated, and in later stages to concentrate on their variation to optimize crystallization conditions. This method has been

26

PATRICIA C. WEBER

implemented using microdialysis cells (Blundell and Johnson, 1976) and used to optimize crystal growth conditions for tryptophan-tRNA ligase (Carter and Carter, 1979) and cytidine deaminase (Betts et al., 1989). An alternative approach, suited particularly to using laboratory robotics, is the successive automated grid search (SAGS) method (Cox and Weber, 1988). The method has been implemented with the hangingdrop crystallization method (McPherson, 1982) and involves the systematic variation of two major crystallization parameters, pH and precipitant concentration, with provisions to vary two others. The variation of solution pH and precipitant concentration effectively varies molecular charge as a function of protein supersaturation in searching for suitable crystallization conditions. The coarse grid is initially used for sampling. Once initial crystals are obtained, the increments of the grid are reduced in the vicinity of the initial successful experiments to optimize crystallization conditions (Fig. 8). The method has been successfully used to crystallize several proteins (Cox and Weber, 1987; Weber et al., 1987). Irrespective of whether crystallization conditions are found by systematic or trial-and-error methods, the process first involves locating some point in the parameter space of possible conditions where the protein will crystallize. Many experiments suggest that the range of parameters over which a given protein will form some sort of crystal is reasonably large (Cox and Weber, 1988; Weber, 1990). In contrast, the parameter space defined by the optimal conditions where crystals suitable for X-ray diffraction studies are grown can be much smaller. Studies of crystal terminal size and the dependence of crystal size on the supersaturation ratio demonstrate that the largest crystals grow within a narrow range of conditions (Heidner, 1978; Ataka and Tanaka, 1986; Betts et al., 1989) (Fig. 7). A typical result is that observed with haloalkane dehalogenase (Rozeboom et al., 1988):Crystals form in ammonium sulfate concentrations greater than 60% saturation over the pH range 5.4-7.2. However, the best crystals form under the more restrictive conditions of 64% saturation and pH 6.3 2 0.1. IN PROTEIN CRYSTALLIZATION XI. NEWDEVELOPMENTS

A . Crystallization in Microgravity

Space exploration offers a unique opportunity to test the effects of gravity on protein crystal growth (Morita, 1985; Bugg, 1986; DeLucas et al., 1986; Drenth et al., 1987). Although it may not be obvious, the growth of millimeter-sized protein crystals in microliter volumes is af-

FIG.8. Approach to protein crystallizat.ion using successive automated grid searches. (A) The bottom rows show the first experiments conducted and typical results using this approach. Three trays each containing 16 droplets are set up. Using the citrate/phosphate buffer system, the pH of the solution is varied from 2.4 to 7.8. A different type of precipitating agent is used in each tray. These are polyethylene glycol (PEG) from a stock solutioti of 25% (w/v) PEG 8000, atnmoniuni sulfate from a saturated stock solution, and a PEG-salt mixture using the 25% PEG solution above and adding 1 M LiCl to all droplets by dilution from a 10 M LiCl stock solution. (Left) The shaded square indicates a crystal-containing droplet. The top two figures in this panel show a typical experimental strategy of expanding and overlapping conditions about those yielding crystals in the firsr wide-screen experiments. (Middle) Occasionally,the distinction between conditions that do and do not cause precipitation is clear, as indicated by shading. In this caSe successive grids explore the region along the precipitation border. (Right) If precipitation occurs in all droplets, the three broad-screen experirncnts are repcated using quite different cotiditions (e.g., higher pH, an alternate temperature or protein concentration, or the addition of cofactors). (R) Successive automated grid searches were applied to the crystallization of a streptavidin-biotin complex. (Top) The initial wide-screen experiment in which the pH of the solution in the columns varied from 2.4 to 7.8 with the citratelphosphate buffer system and the polyethylene glycul (PEG) concentration ranged from 1% to 15% in the row. Droplets were photographed about 2 weeks after beginning the crystallization experiment. Crystals grew overnight at pH 7.8, 15% PEG. These crystals turned brown within 1 week. Crystals that grew more slowly (e+, at pH 6.4) did not discolor even after several months. (Bottom) An expansion of crystallization conditions about those producing crystals it, (A). Magnification: (top) 10.3 x and (hottom) 3.5 x . From Cox and Weber (1987).

FIG.8B. See legend on p. 27.

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

29

fected by gravity in several ways. Protein crystals are more dense than the bulk solution, so that an immediate advantage to crystallization in microgravity is the elimination of crystal sedimentation. Instead of falling to the bottom of the crystallization solution, where fused aggregates can form, crystals grow suspended in solution at the site of nucleation. As a result larger single crystals with more perfect habits are frequently observed in microgravity crystallizations (DeLucas et al., 1986). Density gradients are established at several stages in the crystallization process (Fig. 5). As molecules attach to the growing crystal surface, the solution near the crystal is depleted of solute and becomes less dense than the bulk solution. Under the influence of gravity, such density differences result in convection currents. However, in microgravity, solutions with different densities are not subject to convection, so that solutions mix with less turbulence (Littke and John, 1984) and equilibration between solutions is much slower (DeLucas et al., 1986). Solution turbulence could affect several stages of protein crystal growth. Littke and John (1986) suggested that the rapid onset of P-galactosidase crystallization observed on Spacelab 1 was attributable to a lack of convective mixing turbulence that otherwise disrupted prenuclear molecular complexes in terrestrial control experiments. In terrestrial control experiments isocitrate lyase crystallizes with a dendritic habit (Fig. 4C) that results from local concentration fluctuations at the growing crystal surface, which cause unusually rapid growth along principal crystal axes (Langer, 1989). A more regular habit was obtained in microgravity (Fig. 4D), consistent with the expected elimination of convection-induced concentration fluctuations at the crystal surfaces. Growth in the quiescent microgravity environment may also improve the internal order of protein crystals independent of increases in crystal size (DeLucas et al., 1989). Taken together, the initial data on protein crystal growth suggest that the mechanisms for introduction of lattice defects frequently associated with turbulent growth affect both crystal order and terminal size. Some of the advantages of crystallization in microgravity can be achieved by crystallization in gels (Robert and Lefaucheux, 1988). For example, crystals remain suspended in the gel, although the gel matrix is sufficiently flexible to allow crystal growth. Convection currents are attenuated so that nucleation is reduced and mass transfer occurs primarily by diffusion. Entrapment of foreign particles reduces heterogeneous nucleation, and initial studies suggest that homogeneous nucleation is restricted to the largest pores of the gel containing enough solute molecules to form a critical nucleus. Hen egg white lysozyme and por-

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PATRICIA C . WEBER

cine trypsin have been crystallized in 1% ( v h ) tetramethoxysilane and 0.4% agar gels (Robert and Lefaucheux, 1988). B . Automated Crystallization

Laboratory automation has been adapted for protein crystallization. Setting up a crystallization experiment involves several liquid-handling operations-including dilution, mixing, and dispensing of multicomponent solutions-that are readily automated. Automation reduces manual labor and increases reproducibility by reducing errors and improving the accuracy of solution delivery. Hanging- and sitting-drop crystallizations have been automated to varying extents, ranging from liquid handling for setup (Kelders et al., 1987; Morris et al., 1989) to mixing reservoir solutions from concentrated stocks and combining them in the crystallization droplet (Cox and Weber, 1987) (Fig. 9) to attempts at total automation, in which robotics are additionally used to grease, flip, and seal coverslips on the individual vapor diffusion wells

FIG. 9. An automated system for protein crystallization. This system, composed of a programmable pipetting station, a rotary valve, and a computer, is designed to manipulate liquids automatically for protein crystallization experiments by the hanging-drop technique. Stock solutions are placed at ports of the rotary valve shown at the left. Liquids are dispensed into the wells of the crystallization tray by the 2-ml syringe of the pipetting station. The dispense tip of the pipetting station moves vertically while the crystallization plate moves in the horizontal plane below it. Once the wells are filled, a droplet from each well is placed on the coverslip using the 40-c(1 transfer syringe (shown on the right). A vial of protein solution is placed on the coverslip holder (lower right) after all well solutions have been transferred to the coverslips. In the final step the protein solution is added to the droplets of well solution again using the 4O-pl transfer syringe.

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

Computer

FIG. 10. Components of a data acquisition system for recording the results of crystallization experiments. Sixteen experiments are conducted in a 4 x 4 array on a tray. Each time the crystallization tray is examined visually under the microscope, the bar code label on the tray is first scanned with the bar code reader and a 4 X 4 grid appears on the computer terminal. A rating scheme of 10 descriptive comments is used to evaluate experimental results. Droplet ratings are entered into the computer using the 10 function keys, each corresponding to one of the ratings. When the observation for an individual droplet is entered, the corresponding grid position on the terminal display is filled by a color corresponding to the rating. The grid space is filled with a different color for each rating, rather than a number, to aid in the recognition of input data and to decrease data entry errors. Droplet data, including pH, the concentrations of precipitating agent and additives, and previous ratings, can be displayed for the entire grid by depressing a specified key. During the data entry procedure, the complete description of the tray is also accessible from the database. After the data for all 16 droplets have been entered, they are stored in the computer memory. A hard copy of the ratings is printed on a sticker that can be peeled from the backing and placed in the laboratory notebook. Results of other analyses of the database and the bar codes are printed on the laser printer.

(Ward et al., 1988; Jones et al., 1987). A crystallization plate, on which the protein solution is sandwiched between glass plates, was designed for the automated visual inspection of crystallization experiments (Jones et al., 1987). Photographs of crystals first produced by an automatcd instrument together with some reproduced from the literature 3.'. shown in Fig. 4E-H. As a complement to the automated setup of crystallization experiments, a database system for recording crystallization results (Fig. 10) has been developed to facilitate data acquisition and to aid in the design of subsequent experiments.

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XII. SUMMARY REMARKS

Despite reports of nearly 100 new protein crystal forms in 1988, crystallization remains an important obstacle to many structural studies of biologically important molecules (DeLucas and Bugg, 1987; Giege et al., 1988). Although much remains to be learned, research in this area reveals several recurrent themes, outlined here. 1. Proteins typically appear to be able to form crystals over a fairly large range of solution conditions. However, conditions required to produce large ordered crystals occupy a smaller fraction of the total crystallization parameter space. This observation is important in designing strategies to search for crystal growth conditions. 2. Stochastic events are important in protein crystallization. Solution fluctuations provide the driving force for homogeneous nucleation in supersaturated solutions. Effects are amplified in the small solution volumes used in vapor diffusion experiments, where high surface areavolume ratios produce transient concentration gradients (Fehribach and Rosenberger, 1989). Similarly, temperature and vibration are important factors in controlling crystal nucleation and growth (Feher and Kam, 1985). 3. Solution parameters change during crystal growth. The bulk protein concentration decreases as crystals grow, while the concentration of impurities increases. T h e growing crystals produce concentration gradients in solution. At the same time, electrostatic surface properties of the crystal can alter the activity o f charged components in solution (Rosenberger, 1986; Young et al., 1988). 4. Many crystallization reports emphasize the need to use pure proteins to ensure crystal reproducibility. The application of recombinant DNA technology to the production of truncated gene products promises to alleviate many of the difficulties associated with purifying protein fragments produced with proteolytic enzymes. 5 . Changes in a single experimental parameter can simultaneously influence several aspects of a crystallization experiment. For example, temperature changes affect protein solubility, rates of nucleation and growth, and equilibration of the experimental apparatus. The interaction of parameters makes it difficult to design experiments to isolate individual effects and likewise complicates the interpretation of experimental results. 6. Many lines of evidence suggest that molecular preassociation may be important for protein nucleation and growth.

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

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ACKNOWLEDGMENTS T h e author thanks R. C. DeMattei, R. S. Feigelson, R. Giege, A. McPherson, and D. Ollis for providing manuscript preprints prior to publication and F. R. Salemme for critical discussions.

REFERENCES Althoff, S., Zambrowicz, B., Liang, P., Glaser, M., and Phillips, G. N., Jr. (1988). J. Mol. Biol. 199,665-666. Anderson, W. F., Boodhoo, A., and Mol, C. D. (1988). J. Cryst. Growth 90, 153- 159. Arakawa, T., and Timasheff, S. N. (1984). Bzochemistry 23,5912-5923. Arakawa, T., and Timasheff, S. N. (1985). Biochemistry 24,6756-6762. Ataka, M.. and Asai, M. (1988).J. Cryst. Growth 90, 86-93. Ataka, M., and Tanaka, S. (1986). Biopolymers 25, 337-350. Baker, E. N. (1988).J. Mol. Biol. 203, 1071-1095. Baldwin, E. T., Crumley, K. V., and Carter, C. W., Jr. (1986). [email protected]. 49,47-48. Banerjee, S . K., Pogolotti, A., Jr., and Rupley, J. A. (1975). J. Biol. Chem. 250, 82608266. Betts, L., Frick, L., Wolfenden, R., and Carter, C. W., Jr. (1989). J. Biol. Chem. 264, 6737-6740. Blundell, T. L., and Johnson, L. N. (1976). “Protein Crystallography,” pp. 59-81. Academic Press, New York. Boistelle, R., and Astier, J. P. (1988).J. Cryst. Growth 90, 14-30. Boodhoo, A., Mol, C. D., Lee, J. S., and Anderson, W. F. (1988). J . Biol. Chem. 263, 18578-18581. Bott, R. R., Navia, M. A., and Smith, J. L. (1982).J. Biol. Chem. 257,9883-9886. Brennan, R. G., Vasu, S., Matthews, B. W., and Otsuka, A. J. (1989). J . Biol. Chem. 264,5. Brick, P., Ollis, D., and Steitz, T. A. (1983). J. Mol. Biol. 166, 453-456. Broom, M. €3. H., Witherow, W. K., Snyder, R. S., and Carter, D. C. (19SS).J. Cryst. Growth 90,130-135. Bugg, C. E. (1986).J. Cryst. Growth 76,535-544. Carter, C. W., Jr. (1988).J. Cryst. Growth 90, 168-179. Carter, C. W., Jr., ed. (1990). Methods: Companion Meth. Enzymol. 1. Carter, C. W., Jr., and Carter, C. W. (1979).J. Biol. Chem. 254, 12219-12223. Carter, C. W., Jr., Baldwin, E. T., and Frick, L. (19SS).J. Cryst. Growth 90, 60-73. Carter, D. B., Curry, K. A,, Tomich, C.3. C., Yem, A. W., Deibel, M. R., Tracey, D. E., Paslay, J. W., Carter, J. B., Theriault, N. Y., Harris, P. K. W., Reardon, I. M., ZurcherNeely, H. A,, Heinrikson, R. L., Clancy, L. L., Muchmore, S. W., Watenpaugh, K. D., and Einspahr, H. M. (1988). Proteins: Struct., Funct. Genet. 3, 121-129. Caspar, D. L. D., Clarage, J., Salunke, D. M., and Clarage, M. (1988). Nature (London) 332, 659-662. Cox, M. J., and Weber, P. C. (1987). J. Appl. Crystallop. 20, 366-373. Cox, M. J., and Weber, P. C. (19SS).j. Cryst. Growth 90, 318-324. Cygler, M., Boodhoo, A,, Lee, J. S., and Anderson, W. F. (1987). J . Biol. Chem. 262, 643-648. Davey, R. J. (1986). J. Cryst. Growth 76,637-644. DeLucas, L. J., and Bugg, C. E. (1987). T I B T E C H 5 , 188-193. DeLucas, L. J., Suddath, F. L., Snyder, R., Naumann, R., Broom, M. B., Pusey, M., Yost,

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V., Herren, B., Carter, D., Nelson, B., Meehan, E. J., McPherson, A., and Bugg, C. E. (1986).J. Cyst. Growth 76,681-693. DeLucas, L. J., Smith, C. D., Smith, H. W., Senadhi, V., Sendhi, S. E., Ealick, S. E., Carter, D. C., Snyder, R.S., Weber, P. C., Salemme, F. R.,Ohlendorf, D. H., Einspahr, H. M., Clancy, L. L., Navia, M. A., McKeever, B. M., Nagabhushan, T. L., Nelson, G., McPherson, A., Koszelak, S., Taylor, G., Stammers, D., Powell, K., Darby, G., and Bugg, C. E. (1989). Science 246,651-654. DeMattei, R. C., and Feigelson, R. S. (1989).J . Cryst. Growth 97,333-336. Dijkstra, B. W., Drenth, J., Kalk, K. H., and Vandermaelen, P. J. (1978).J. Mol. Biol. 124, 53-60. Drenth, J., Helliwell, J. R., and Littke, W. (1987). In “Fluid Sciences and Materials Science in Space” (H. U. Walter, ed.), pp. 451-476. Springer-Verlag, Berlin and New York. Dreusicke, D., Karplus, P. A., and Schulz, G. E. (1988).J . Mol. Bzol. 199, 359-371. Durbin, S. D., and Feher, G. (1986).J . Cryst. Growth 76,583-592. Edwards, S. L., Kraut, J., Xuong, N., Ashford, V., Halloran, T. P., and Mills, S. E. (1988). J . Mol. Biol. 203,523-524. Feher, G., and Kam, Z. (1985). In “Methods in Enzymology” (H. W. Wyckoff, C. H. W. Hirs, and S. N. Timasheff, eds.), Vol. 114, pp. 77-1 12. Academic Press, Orlando, Florida. Fehribach, J. D., and Rosenberger, F. (1989).J. Cryst. Growth 94,6- 14. Feigelson, R. S. (1988).J . Cryst. Growth 90, 1 - 13. Fiddis, R. W., Longman, R. A., and Calvert, P. D. (1979).J. Chem. Soc., Furaday Tram. 75, 2753-2761. Finzel, B. C., and Salemme, F. R. (1985). Nature (London) 315,686-688. Finzel, B. C., and Salemme, F. R.(1986). Biophys.J. 49,73-76. Fitzgerald, P. M. D., and Madsen, N. B. (1986).J. Cryst. Growth 76,600-606. Fowlis, W. W., DeLucas, L. J., Twigg, P. J.. Howard, S. B., Meehan, E. J., Jr., and Baird, J. K. (1988).J. Cryst. Growth90, 117-129. Frey, M., Genovesio-Taverne, J.-C., and Fontecilla-Camps,J. C. (1988).J. Cryst. Growth 90, 245-258. Furuno, T., and Sasabe, H. (1985). Kzno Zazryo 5,77-81. Garavito, R. M., Markovic-Housley, Z., and Jenkins, J. A. (1986). J . Cryst. Growth 76, 701 -709. Gernert, K. M., Smith, R.,and Carter, D. C. (1988). Anal. Biochem. 168, 141 -147. Giege, R., Moras, D., Ruff, M., Thierry, J. C., Hirsch, E., Rodeau, J. L., and Mikol, V. (1988). Proc. Int. Biotechnol. Symp., 8th pp. 1 - 1 1. Gilliland, G. L. (1988).J . Cryst. Growth 90,5 1-59. Gilliland, G. L., and Bickham, D. (1990). Methods: Companion Meth. Enrymol. 1,6- 1 1 . Gilliland, G. L., and Davies, D. R. (1984). In “Methods in Enzymology” (W. B. Jakoby, ed.), Vol. 104, pp. 370-381. Academic Press, Orlando, Florida. Gilliland, G. L., Winborne, E. L., Masui, Y., and Hirai, Y. (1987). J. Biol. Chem. 262, 12323- 12324. Harburn, G., Taylor, C. A., and Welberry, T. R. (1975). “Atlas of Optical Transforms.” Cornell Univ. Press, Ithaca, New York. Heidner, E. (1978).J. Cryst. Growth 44, 139- 144. Hirsch, R. E., Lin, M. J., and Nagel, R.L. (1988).J . Biol. Chem. 263,5936-5939. Hofrichter, J., Ross, P. D., and Eaton, W. A. (1974). Proc. Natl. Acud. Scz. U.S.A.71, 4864-4868. Hofrichter, J., Ross, P. D., and Eaton, W. A. (1976). Proc. Natl. Acad. Sci. U.S.A. 73, 3035-3039.

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

35

Jacoby, W. B. (1968). Anal. Biochem. 26,295-298. Jones, N. D., Decter, J. B., Swartzenderber, J. K., and Landis, P. L. (1987). Am. c r y s t d o f f . Assoc. Meet., March 15-20, Austin, Texas Abstr. H-4. Kam, Z., Shore, H. B., and Feher, G. (1978).J. Mol. Bzol. 123,539-555. Kelders, H. A., Kalk, K. H., Gros, P., and Hol, W. G. J. (1987). Protein Eng. 1, 301-303. Langer, J. S. (1989). SciaCe243,1150-1156. Leung, C. J.. Nall, B. T., and Brayer, G. D. (1989).J. Mol. B i d . 206, 783-785. Lippmann, C., Betzel, C., Dauter, Z., Wilson, K., and Erdmann, V. A. (1988). FEBS Lett. 240,139-142. Littke, W., and John, C. (1984). Science 225, 203-204. Littke, W., and John, C. (1986).J. Cryst. Growth 76, 663-672. Lorber, B., Kern, D., Mejdoub, H., Boulanger, Y., Reinbolt, J., and Giege, R. (1987). Eur. J . Biochem. 165,409-417. Matsuura, Y. (1985). Nippon Kessho Seicho Gakkaishi 12, 106-1 16. McPherson, A. (1982). “The Preparation and Analysis of Protein Crystals.” Wiley, New York. McPherson, A. (1990). Eur.J. Biochem. 189, 1-23. McPherson, A,, and Shlichta, P. (1988). Science 239,385-387. McPherson, A,, and Spencer, R. (1975). Arch. Biochem. Biophys. 169,650-661. Michel, H. (1983). Trends Biochem. Sci. (Pers. Ed.) 8,56-59. Mikol, V., Rodeau, J.-L., and Giege, R. (1989).J. Appl. Cystallogr. 22, 155-161. Morita, Y. (1985). Pharm. Tech.Jp,. 1,991-995. Morris, D. W., Kim, C. Y., and McPherson, A. (1989). Biotechniques 7,522-527. Ollis, D., and White, S. (1990). In “Methods in Enzymology” (M. P. Deutscher, ed.), Vol. 182, pp. 646-659. Ollis, D. L., Brick, P., Abdel-Meguid, S. S., Murthy, K., Chase, J. W., and Steitz, T. A. (1983).J. Mol. Biol. 170,797-801. Orbell, J. D., Guddat, L. W., Machin, K. J., and Isaacs, N. W. (1988). Anal. Biochem. 170, 390-392. Pahler, A., Hendrickson, W. A,, Gawinowicz Kolks, M. A., Argarana, C. E., and Cantor, C. E. (1987).J. Biol. Chem. 262, 13933-13937. Perry, A. L., and Palmer, R. A. (1988).J. MoZ. Bzol. 201,659-660. Prasad, L., Vandonselaar, M., Lee, J. S., and Delbaere, L. T. J. (1988). J. B i d . Chem. 263, 2571-2574. Pusey, M. L., Snyder, R. S., and Naumann, R. (1986).J. B i d . Chem. 261,6524-6529. Pusey, M., Witherow, W., and Naumann, R. (1988).J. Cryst. Growth 90, 105- 11 1. Robert, M. C., and Lefaucheux, F. (1988).J. C y s t . Growth 90, 358-367. Rose, J. P., Yang, D., Yoo, C. S., Sax, M., Breslow, E., and Wang, B.-C. (1988). Eur. J. Biochem. 174, 145-147. Rosenberger, F. (1986).J. Cyst. Growth 76,618-636. Rosenberger, F., and Meehan, E. J. (1988).J. Cryst. Growth 90,74-78. Rozeboom, H. J., Kingma, J., Janssen, D. B., and Dijkstra, B. W. (1988). J . Mol. B i d . 200, 61 1-612. Ruff, M., Cavarelli, J., Mikol, V., Lorber, B., Mitschler, A., Giege, R., Thierry, J. C., and Moras, D. (1988).J. Mol. B i d . 201, 235-236. Salemme, F. R. (1972). Arch. Biochem. Biophys. 151,533-539. Salemme, F. R., Genieser, L., Finzel, B. C., Hilmer, R. M., and Wendoloski, J. J. (1988). J. Cryit. Growth 90,273-282. Schlichtkrull, J. (1957). Acta Chem. Scand. 11,439-460.

36

PATRICIA C . WEBER

Sheriff, S., Hendrickson, W. A., Stenkamp, R. E., Sieker, L. C., and Jensen, L. H. (1985). 82, 1104- 1107. Proc. Natl. Acad. Sci. U.S.A. Sheriff, S., Hendrickson, W. A., and Smith, J. L. (1987). J. Mol. Biol. 197,273-296. Sheshadri, B. S., and Venkatappa, M. P. (1983).J . Sci. Ind. Res. 42,615-627. Spangler, B. D., and Westbrook, E. M. (1989). Biochemistry 28, 1333- 1340. Steitz, T. A. (1971).J. Mol. Biol. 61,695-700. Stura, E. A., and Wilson, I. A. (1990). Methods: Companion Meth. Enzymol. 1,38-49. Takizawa, T., and Hayashi, S. (1976).J . Phys. SOC.Jpn. 40,299-300. Thaller, C., Weaver, L. H., Eichele, G., Wilson, E., Karlsson, R.,and Jansonius, J. N. (1981).J.Mol. Biol. 147,465-469. Timasheff, S. N., and Arakawa, T. (1988).J. Cryst. Growth 90,39-46. Van Der Laan, J. M., Swarte, M. B. A., Groendijk, H., Hol, W. G. J., and Drenth, J. (1989). Eur. J . Biochem. 179,715-724. Ward, K. B., Perozzo, M. A., and Zuk, W. M. (1988). J. Cryst. Growth 90,325-339. Weber, P. C. (1990). Methods: Companion Meth. Enzymol. 1, 31-37. Weber, P. C., and Salemme, F. R. (1977).J. Mol. Biol. 117, 815-820. Weber, P. C., Cox,M. J., Salemme, F. R., and Ohlendorf, D. H. (1987).J . Biol. Chem. 262, 12728-12729. Wyckoff, H. W., Hirs, C . H. W., and Timasheff, S. N. (eds.). (1985). “Methods in Enzymology,” Vol. 114. Academic Press, Orlando, Florida. Yonath, A., Mussig, J., and Wittmann, H. G. (1982). J . CellBiochem. 19, 145-155. Young, C. C., DeMattei, R. C., Feigelson, R. S., and Tiller, W. A. (1988). J . Cryst. Growth 90,79-85.

PROTEIN HYDRATION AND FUNCTION By JOHN A. RUPLEY* and GlORGlO CARERlt 'Department of Biochemistry. University of Arizona. Tucson. Arizona 85716 tDlpartimento di Flsica. Universita di Roma I. Rome 00185. Italy

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. Thermodynamics . . . . . . . . . . . . . . . . . . . . . . . . . . .

A. Powders and Films . . . . . . . . . . . . . . . . . . . . . . . . B. Denaturation . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Nonfreezing Water . . . . . . . . . . . . . . . . . . . . . . . . D. Solutions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I11. Dynamics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Dielectric Relaxation . . . . . . . . . . . . . . . . . . . . . . . . B. Percolation Model . . . . . . . . . . . . . . . . . . . . . . . . . C. Resonance . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Hydrogen Exchange . . . . . . . . . . . . . . . . . . . . . . . . E . Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Enzyme Activity . . . . . . . . . . . . . . . . . . . . . . . . . . G. Other Measurements . . . . . . . . . . . . . . . . . . . . . . . IV. Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A . Diffraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Computer Simulation . . . . . . . . . . . . . . . . . . . . . . . . . A. Molecular Dynamics . . . . . . . . . . . . . . . . . . . . . . . . B . Monte Carlo Simulations . . . . . . . . . . . . . . . . . . . . . . C . Accessible Surface andThermodynamics of Hydration . . . . . . . . D. Other Simulations . . . . . . . . . . . . . . . . . . . . . . . . . VI . Picture of Protein Hydration . . . . . . . . . . . . . . . . . . . . . . A. Fully Hydrated Protein . . . . . . . . . . . . . . . . . . . . . . B . Hydration Process . . . . . . . . . . . . . . . . . . . . . . . . . C. Bound Water . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Hydration and Conformation . . . . . . . . . . . . . . . . . . . . VII . Hydration and Function . . . . . . . . . . . . . . . . . . . . . . . . A.Folding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Chemistry of Transition States . . . . . . . . . . . . . . . . . . . C. Protonic Conduction and Percolation . . . . . . . . . . . . . . . . D. Substrate Binding . . . . . . . . . . . . . . . . . . . . . . . . . E . Water Networks . . . . . . . . . . . . . . . . . . . . . . . . . . F. Fluctuations and Protein Motions . . . . . . . . . . . . . . . . . . G. Fluctuations and Catalysis . . . . . . . . . . . . . . . . . . . . . H . Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . I . Complex Systems . . . . . . . . . . . . . . . . . . . . . . . . . VIII . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix: Percolation Theory . . . . . . . . . . . . . . . . . . . . . A. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . .

38 41 41 52 54 56 61 61 69 71 80 84 91 95 99 99 107 112 112 115 117 120 122 126 131 137 139 141 142 143 145 146 147 148 148 149 150 152 154 154

37 ADVANCES IN PROTEIN CHEMISTRY. Vol. 41

Copyright 0 1991 by Academic Press. Inc. All rights of reproduction in any form reserved.

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JOHN A. RUPLEY AND GIORGIO CARER1

B. Invariant Quantities . . . . . . . . . . . . . . . . . . . . . . . . C. Finite-Size Effects . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

156 160 161

I . INTRODUCTION In 1974 Kuntz and Kauzmann reviewed the hydration of proteins for Advances in Protein Chemistry. Their work covered the field critically and thoroughly, stimulated wide interest, and laid a foundation for the now 5-fold greater number of publications per year on this subject. It no longer appears appropriate to attempt so comprehensive a review. This article covers topics that bear most directly on descriptions of the hydration shell and the hydration process. Questions that were open in 19’74, such as the extent of the interface between protein and solvent and the degree to which the properties of the interface differ from the bulk phase, are now largely resolved. There are, of course, new open questions, many of which concern the way in which the hydration shell participates in protein function. The existence of a hydration shell was recognized early. Hydrodynamic measurements that were used to define the overall size and shape of proteins were used also in analyses aimed at delimiting the amount of solvent carried along by them (Edsall, 1953; Scheraga and Mandelkern, 1953). Most physical techniques that serve to describe proteins have been applied also to description of their interface with solvent. Table I lists the principal methods for investigating protein hydration and shows the type of information obtained-specifically, whether the measurement gives information primarily on protein or solvent properties, on structure, on time-average or dynamic processes, or on powder or solution samples. This review focuses on work with partially hydrated powder samples. To understand the interaction of water at the protein surface, the water activity should be varied over a wide range, as is possible for powder samples hydrated in a controlled atmosphere, but not generally for solutions. Hydration can be considered a process, that of adding water incrementally to dry protein, until a level of hydration is reached beyond which further addition of water produces no change and only dilutes the protein. The hydration process has several stages and an end point, reflected similarly in different types of measurements. Time-average measurements appear to have a common pattern and most closely fit a single picture of the process. Dynamic measurements sometimes show,

TABLE I Methodsfor Measurement of Hydrationn State of sample Measurement Diffraction X-Ray, neutron Spectroscopy IR, UV-VIS, circular dichroism Neutron scattering, Mossbauer Spectroscopy and relaxation NMR, ESR Dielectric Solution thermodynamics C,, V, preferential binding, hydration forces Sorption thermodynamics G , H, C, V Hydrodynamics Sedimentation, viscosity Computer simulation Molecular dynamics and Monte Carlo, surface, and packing simulations Special rate properties Enzyme activity, hydrogen exchange Special time-average properties Nonfreezing water Unfolding transition

Solution

+ + + (+)

Powder/film Time-average

(+)

+ + +

+ t

+

+ + + +

(+)

+ + +

nAdapted from Table I of Rupley et al. (1983).This list is not exhaustive.

Dynamic

Structural

Water

Macromolecule

+

+

+

+ +

+ + (+)

(+)

+

+

+

+

(+)

+ +

Information about

Type of information

+ +

+

+ (+)

+

40

JOHN A. RUPLEY AND GIORGIO CARER1

often understandably, a dependence on hydration level that lies outside the common pattern of the time-average results. The hydration shell can be defined as the water associated with the protein at the hydration end point. This shell represents monolayer coverage of the protein surface. Water outside the monolayer is perturbed to a significantly smaller extent, typically not detected by measurements of properties such as heat capacity, volume, or heat content. Thus, in a concentrated protein solution or in a wet protein powder, at hydration levels above the hydration end point, the perturbation of multilayer solvent is small compared with the stronger perturbation within the monolayer, and the former can be viewed as a second-order effect. However, the cumulative contribution from small perturbations can be detected in certain time-average measurements made with sufficiently large aggregates of proteins, nucleic acids, or lipids-for example, measurements of the hydration forces described by Parsegian and Rand (Parsegian et al., 1986; Rand et al., 1985) and by Israelachvili and Marra (1986). Dynamic properties that exhibit a first-order effect of hydration beyond the monolayer are best considered case by case. This review has three major parts. The first, comprising Sections II-V, is a summary of the literature that bears on the protein hydration process and the hydration shell, categorized by type of measurement. It is assumed that the reader has access to the reviews by Kuntz and Kauzmann (1974) and by Edsall and McKenzie (1983), and only the most central of the results described by them are described again here. We hope that our predilection for some types of measurement does not produce a distorted image of hydration. In this regard the reader is referred in Sections II-V to recent reviews, which provide fuller coverage of topics that are treated incompletely here. The second and third parts of this review develop, through correlation of the results described in the first part, a picture of the hydration process and the hydration shell (Section VI) and an assessment of how the hydration shell may modulate enzyme and other protein functions (Section VII). The literature on protein hydration is now rich enough to provide a comfortably detailed picture of the protein-water interface. The ways in which the interface enters into function are just beginning to emerge, and one purpose here is to point out directions in which one may move to understand better the relationship to function. Sections VI and VII are meant to stand alone as summary statements, and some overlap with the preceding sections describing results should be expected. Several studies have shown how proton movement on a lightly hydrated protein or in water-poor regions of other systems can be under-

PROTEIN HYDRATION AND FUNCTION

41

stood by using the percolation model. This model applies to stochastic processes that evolve on a topologically disordered matrix. It focuses on the long-range connectivity established at a threshold coverage of a surface or volume with conducting or otherwise functional elements. T h e explosive growth of a percolative process above the threshold level is characteristic of a phase transition. The region near the percolation threshold is similar to the critical regions of more familiar phase transitions. Percolative behavior, only recently demonstrated for a protein system, is not generally known to biochemists and biologists. Because percolation should come to be more widely recognized as having a role in biology, a tutorial Appendix is included as an introduction to percolation theory. T h e attention of the reader is drawn to several books and reviews on protein hydration, in addition to the reviews by Kuntz and Kauzmann (1974) and by Edsall and McKenzie (1983). Recent volumes of Method in Enzymology (Hirs and Timasheff, 1985; Packer, 1986) describe measurements on the hydration of protein and membrane systems. Saenger ( 1987) has reviewed aspects of macromolecule hydration. Edsall ( 1980) has given a brief history of research on water. Several summaries of current research in biophysics describe work related to the hydration of macromolecules (Clementi and Chin, 1986; Ehrenberg et al., 1987; Moras et al., 1987; Welch, 1986). For comprehensive treatments of the properties of water and aqueous solutions, see the multivolume treatise by Franks (e.g., Franks, 1979), the review by Edsall and McKenzie (1978), and the volume by Eisenberg and Kauzmann (1969). 11. THERMODYNAMICS

A . Powders and Films 1 . Sorption Sorption isotherms, measurements of the weight of water adsorbed by a protein sample as a function of the partial pressure of water in the vapor phase at constant temperature, are among the earliest descriptions of protein hydration [for references to early work, see Bull and Breese (1968a) and McLaren and Rowen (1951)l. A typical isotherm, for lysozyme, is shown in Fig. 1. There is a “knee” at 0.05-0.1 h (g of water per g of protein, mass ratio) and a strong upswing to the isotherm near 0.25 h. Sorption measurements on model polymers and chemically modified proteins (see Watt and D’Arcy, 1976;

42

JOHN A. RUPLEY AND GIORGIO CARER1

I

I

I

I

0.

0. h

0

0.50 PIP0

0.75

I .oo

FIG. 1. D 2 0 sorption isotherm for lysozyme at 27°C. The data were fit by a model with three classes of sorption sites, to give curves (a), (b), and (c). From Careri et al. (1979b).

Rochester and Westerman, 1976a,b, 1977; and references cited therein) suggested the molecular basis for the sigmoidal shape of the isotherm. Below the knee water interacts principally with ionizable protein groups. In the plateau region, between 0.1 and 0.25 h, water binds to polar sites. Above 0.25 h water condenses onto the weakest binding sites of the protein surface to complete the hydration process, and at sufficiently high water content (water partial pressure) the system passes into the solution state. Methods of measurement have been reviewed (Kuntz and Kauzmann, 1974; McLaren and Rowen, 1951; Poole and Finney, 1986). Hydration levels are often established by isopiestic equilibration of protein samples against concentrated salt or sulfuric acid solutions of known water vapor pressure. A difficulty with this method is the long equilibration time, possibly several days, which is likely a consequence of the sample size (typically 100 mg or larger). Wilkinson et al. (1976) have described an automated sorption isotherm device; transducers are used for the measurement of vapor pressure and sample weight. Gascoyne and Pethig (1977) used a resonating quartz crystal microbalance to study the hydration of bovine serum albumin and other proteins. Rao and Bryan (19'78)

PROTEIN HYDRATION AND FUNCTION

43

and Sherman et al. (1973) used Karl Fischer titration to determine small amounts of tightly held water. Gevorkyan and Morozov (1983) measured sample weight by the vibrational behavior of a needle to which the sample was attached. The mass ratio h is commonly used in describing protein hydration. Hydration, however, should depend more closely on protein surface than on volume or mass. Most of the data described in this review are for small globular proteins, for which weight and surface-based measures should be similar. Comparisons of proteins of much different size may need to take into account surface area, compactness, and domain size. Theoretical treatments of the sorption process for proteins, generally undertaken with the intent of explaining the sigmoidal shape typical of protein isotherms, were reviewed by Kuntz and Kauzmann (1974). Most interpretations are based on the Brunauer-Emmett-Teller (BET) theory (Brunauer et al., 1938) or an extension of it, for which the model is a set of strong-binding surface sites and one or more sets of weaker binding sites (Gascoyne and Pethig, 1977; Kuntz and Kauzmann, 1974). The BET theory and its extensions are analyses of the interaction between surface and vapor; the solution phase is not explicitly introduced into the treatment. From the statistics of polymer solutions, Flory (1953) obtained an isotherm of particularly simple form, with one variable parameter, that describes well the high hydration range of the sorption process and the solution state. Doster et al. (1986) applied Flory’s model to myoglobin. Using scanning calorimetry, they measured the melting point depression, so determining the solvent activity as a function of volume fraction myoglobin in solution. The data from 0 to 0.62 volume fraction could be fit exactly by a Flory-Huggins equation, modified to allow for not all of the segments being flexible. For myoglobin the number flexible was estimated as 40-50 of the 153 residues. Under this assumption the interaction parameter agrees with an experimental determination of the enthalpy of mixing water and hydrated myoglobin. For volume fractions higher than 0.62 the Flory-Huggins equation does not fit the data, which was expected, since this volume fraction corresponds to the point of full hydration of myoglobin: 0.39 h. These experiments were performed in parallel with heat capacity (Section 11) and infrared spectroscopic studies (Section IV) of a low-temperature glass transition, seen also in Mossbauer spectroscopy (Section 111) and other properties (Section VI). Hill (1949) extended the statistical thermodynamic treatment of the Langmuir isotherm to model localized unimolecular adsorption on a random heterogeneous surface, including the case of an adsorbate such

44

JOHN A. RUPLEY AND GIORGIO CARER1

as water, for which there are lateral interactions between adsorbed molecules. This description seems particularly well suited to a protein, where water sites are likely to exhibit a wide range of interaction energies, and for which sites of similar chemistry are more or less randomly distributed about the protein surface. The isotherm can be evaluated by numerical methods. Four variable parameters are enough for a good fit of the isotherm to protein data, including temperature dependence of the sorption. The Hill (1949) analysis predicts two phase transitions within the hydration process: at low hydration (at the knee of the isotherm) there is a two-dimensional condensation of isolated water molecules, distributed about the surface, into clusters; at high hydration (at the end of the plateau region) there is a second condensation over the weakest sites to complete the sorption process. Protein isotherms typically exhibit hysteresis, for which the sorption limb may lie 0.01-0.02 h below the desorption limb. This phenomenon has an important consequence. Conversions of the isotherm information into free energies of transfer of water to the protein surface, and of the temperature dependence of the isotherm into enthalpies of transfer, are done under the assumption that the isotherm reflects an equilibrium state. If there is hysteresis, the equilibrium state may not be defined well enough for thermodynamic analysis. Sorption hysteresis is generally reproducible. Repeated cycling of a single sample can change the response (Luescher-Mattli and Ruegg, 1982a; Rao and Das, 1968). Several explanations of the hysteresis have been offered: (1) capillary condensation, in interstices of the solid material (McLaren and Kirwan, 1976); (2) metastable states associated with phase changes within the adsorbate and corresponding to loci on van der Waals-type loops (Hill, 1947); and (3) conformational changes within the protein, associated with low hydration, that reverse slowly compared with the rate of water sorption (Bryan, 1980; Cerofolini and Cerofolini, 1980; Luescher-Mattli and Ruegg, 1982a).Bryan (1987) has described several possible models. To the above suggestions may be added another: that the number of nucleation sites for condensation of water on the surface is limiting. Charged sites on the protein surface serve as nuclei for condensation of water. If charged sites were absent, condensation should be impossible for a particle as small as a protein (Rupley et d,1983). Thus, in the plateau region of the isotherm, where hysteresis is observed, the addition of water to the surface in a sorption process would be only by addition to those clusters already present and centered on the charged sites. In a desorption process nucleation would not be limiting, and true thermodynamic equilibrium could be reached at all stages of the process, with the number of clusters allowed to exceed the number of charged sites.

PROTEIN HYDRATION AND FUNCTION

45

It is possible, and perhaps generally believed, that the high reproducibility of an isotherm justifies the extraction of thermodynamic values from data that show hysteresis. However, hysteresis would still be a source of systematic error in the values. There is a poorly documented impression that small samples or thin films display less hysteresis. Hysteresis was not found for the heat capacity isotherm (Yang and Rupley, 1979), which may be taken as support for the view that meaningful freeenergy information also can be derived from sorption isotherms. Morozov et al. (1988) have made parallel measurements of the elastic properties (Young’s modulus) and sorption isotherms of monoclinic, triclinic, and tetragonal lysozyme crystals, cross-linked with glutaraldehyde. They estimate that the free energy of deformation is nearly as large as the total free energy of dehydration; that is, it is a principal factor in determining the free energy of the sorption process. The withdrawal of water from a crystal was correlated with its deformation, and this correlation differed among crystal forms. The authors suggested that there is a large contribution from mechanical deformation to the sorption process for protein films and powders also. If true, then the generally accepted methods for extracting thermodynamic data from sorption isotherms lead to incorrect conclusions and incorrect thermodynamic values. This suggestion is interesting. However, it conflicts with other data, for example, measurements of the partial specific volume (discussed below): several protein systems behave ideally to 0.2 h, which would not be observed if there were pores only partially filled with solvent, as in the Morozov model. 2. Enthalfi

T h e temperature dependence of the sorption isotherm defines, through the van? Hoff relationship, the isotherm for the heat of sorption, generally calculated as the isosteric heat, which is the change in partial molar heat content for transfer of water from the vapor phase to the protein. Kuntz and Kauzmann (1974) gave a summary of the thermodynamic functions for sorption processes. The isosteric heat varies strongly with the hydration level. Luescher and Ruegg and collaborators (Luescher and Ruegg, 1978; Luescher et al., 1978, 1979; LuescherMattli and Ruegg, 1982b) analyzed the temperature dependence of the sorption isotherms for a-chymotrypsin and its tosyl derivative. The isosteric heats are large at low h, with an extremum in the heat at the knee of the isotherm. Studies of lysozyme (Hnojewyj and Reyerson, 1961) and ribonuclease (Foss and Reyerson, 1958) also showed an extremum in the differential heat at low coverage of the surface. At half-coverage the differential heat approaches the heat of condensation of pure water.

46

JOHN A. RUPLEY AND GIORGIO CARER1

Berlin et al. (1970) and Ginzburg (1982) used differential scanning calorimetry to measure the heat of desorption of samples of varied initial water content, for several proteins. The data show considerable scatter, probably owing to broad endotherms that span about 100°C. Almog and Schrier (1978) made a direct calorimetric measurement of the dependence of the heat of solution of ribonuclease A on water content (Fig. 2). The heat of solution drops strongly in the low hydration range: 90% of the heat change is obtained at about half-hydration. The differential heat for transfer of water from the pure liquid to the protein is estimated from the data of Fig. 2 as 8 kcal/mol of water at the lowest hydration studied (the heat of condensation of water should be added for comparison with isosteric heats), and it decreases monotonically with increased hydration. There is no extremum at low hydration, unlike what has been reported based on the temperature dependence of the sorption isotherm. It is not clear whether this difference reflects inaccuracies in the data used in van’t Hoff analyses of the sorption isotherms, or a complex hydration path that is not modeled properly in the van’t Hoff analyses. 400

I

300 .

$

m

200 -

c:

I 6 I

100.

20

40

60

80

100

120

140

160

180

200

MOLES OF H 20 PER MOLE OF RIBONUCLEASE A

FIG. 2. The dependence of the negative enthalpy of solution (soh) of lyophilized solid ribonuclease A on the ratio of the number of moles of water contained in the lyophilized solid to the number of moles of ribonuclease A. From Almog and Schrier (1978).

PROTEIN HYDRATION AND FUNCTION

47

3 . Heat Capacity

The heat capacity isotherm is likely to be particularly informative: (1) It can be measured over the full range of system composition, from dry protein to the dilute solution, and thus serves to link studies of powders and solutions. (2) The heat capacity is sensitive to changes in the chemistry of water, including interaction with surface hydrophobic groups, and should sense all time-average events associated with hydration. Using a drop calorimeter, Yang and Rupley (1979) measured the heat capacity isotherm for lysozyme at 25°C (Fig. 3). There is no discontinuity where the system changes from homogeneous solution to wet powder near 0.2 weight fraction protein ( w 2 ) . The linear response between w2 = 0 and w2 = 0.73 requires that the partial specific heat capacities of solvent and protein be invariant in this region, which is equivalent to the only change within this region being the removal of bulk solvent from the system. The break in the linear response at w2 = 0.73 defines the point of full hydration. The irregular response between w2 = 0.73 and wp = 1.0 describes the heat capacity changes associated with hydration of the protein. The point of full hydration determined by the heat capacity response corresponds to 0.38 h, or 300 mol of water per mol of lysozyme. The

WEIGHT FRACTION

FIG. 3. Specific heat of the lysozyme-water system from 0 to 1.0 weight fraction of protein. Least-squaresanalysis of the linear portion of the heat capacity function from 0 to 0.73 weight fraction of water gives Ei2 = 1.483 5 0.009 J K-' g-I. The value of c g = 1.26 2 0.01 J K-' g-'. From Yang and Rupley (1979).

48

JOHN A. RUPLEY AND GIORGIO CARER1

surface area of lysozyme, calculated from the crystal structure (Lee and Richards, 1971; Shrake and Rupley, 1973), is 6000 The average coverage of the surface is thus 20 A2 per water molecule. This is the maximum coverage that can be obtained with a network of water molecules, arranged as in the planes of molecules orthogonal to the c axis of ice I. Thus, the point of full hydration of lysozyme, at 300 mol/mol, cannot correspond to more than one monolayer. Apparently, this monolayer of surface water meshes smoothly with the surrounding bulk water, and the perturbation of the solvent by the protein surface is limited to a one-molecule-thick interface region. This statement is true for timeaverage quantities, to a resolution corresponding to perhaps 100 cal/mol of water. Some thermodynamic measurements, such as the osmotic pressure response, reflect longer-range and weaker perturbations. Figure 4 gives the apparent specific heat capacity [+(cp2)]of lysozyme

w2.

calculated from the data of Fig. 3, as a function of system composition for the water-poor region of the isotherm. The apparent specific heat is the nonideal, or excess, heat capacity, normalized to unit weight of protein. The isotherm can be broken into four regions. 1. R e p o n I, dilute protein solution to 0.38 h: The partial specific heats are constant and the protein is fully hydrated. 1.6 1

1 I

I

I

I

-

I.

I

II

I

I

I

1

I I

0

0.05

0.10

0.15

0.20

0.25

0.30

0.35

,

0.40

0.45

0.50

g WATEWg PROTEIN

FIG.4. The apparent specific heat capacity of lysozyme from 0 to 0.45 g of water per gram of protein. The curve is calculated. The heat capacity measurements were made with lyophilized powders of lysozyme, appropriately hydrated, except for the four measurements indicated by the square symbols, for which the sample was a film formed by slowly drying a concentrated solution of lysozyme. From Yang and Rupley (1979).

PROTEIN HYDRATION AND FUNCTION

49

2. Regaon 11, 0.38 to 0.27 h: Water condenses over the surface elements that interact most weakly with water, to complete the solvent shell. There is a rise and a fall in the heat capacity at the junction of regions I1 and 111, characteristic of an order-disorder transition, which can be understood in terms of the high-coverage phase change described by Hill (1949). 3. Regaon 111,0.27 to 0.07 h: The nonideality of the specific heat decreases to near zero. The partial specific heat content of the adsorbed water is large, being greater than that of the pure liquid. 4. Regzon IV, 0.07 h to d v protein: There is a reaction heat contribution, centered at 0.05 h, that other measurements (Careri et al., 1979b) have shown to be the result of normalization of the pK order for the lightly hydrated protein (transfer of protons from carboxylate to other protonatable groups). The partial specific heat of the hydration water is 3.3 J K - ' g-l in region IV and 5.8 J K-l g-l in region 111. This difference is consistent with a transition between a state in which the adsorbed water is predominantly dispersed to one in which there are hydrogenbonded clusters. This transition corresponds to the two-dimensional condensation at low coverage of the surface predicted by the Hill (1949) model. The region IV-region I11 transition of the heat capacity isotherm corresponds to the knee of the sorption isotherm, and the region IIIregion I1 transition corresponds to the strong upswing in the sorption isotherm. The heat capacity measurements for lysozyme are consistent with data obtained for other globular protein systems, for example, ovalbumin (Bull and Breese, 1968b; Suurkuusk, 1974), chymotrypsinogen (Hutchens et al., 1969; Suurkuusk, 1974), and insulin (Hutchens et al., 1969). For a discussion see Yang and Rupley (1979). Several measurements have been made of the low-temperature heat capacities of proteins. 1. Hutchens et al. (1969) determined the heat capacities of zinc insulin at 0 and 0.04 h and of chymotrypsinogen A at 0 and 0.107 h, from 10 to 310 K. For all samples the data were a smooth function of temperature, with no indication of a glass or phase transition at any temperature. The absence of a phase transition corresponding to the iceliquid water transition is expected for low hydrations. These appear to be the only data in the literature that have been used to determine the entropy of a protein sample. Hutchens et al. (1969) calculated the standard entropy of formation of a peptide bond as 9.0-9.3 cal K-l mol-l. 2. Doster et al. (1986) determined the specific heat of water in myoglobin crystals from 180 to 270 K, as the difference between scanning

50

JOHN A. RUPLEY AND GIORGIO CARER1

calorimetric measurements made on wet myoglobin crystals and the same sample after drying. The crystal water showed a glass temperature near 220 K. Infrared spectroscopic measurements were carried out in parallel (Section IV). 3. Andronikashvili et al. (1979) measured the heat capacity of collagen at 0 and 0.4 h, from 4 to 320 K. Neither sample showed an iceliquid water phase transition. The anhydrous sample showed a smooth increase in heat capacity with temperature. The hydrated sample showed a discontinuity at 120 K, apparently associated with an order-disorder transition, perhaps a glass transition, above which there was a 10-fold stronger dependence of the heat capacity on temperature. 4. Haly and Snaith (1968) measured the specific heat of the waterwool system from 0 to near 0.4 h, for temperatures from - 70 to 100°C. At high temperature (i.e., 80°C) the discontinuity between regions I11 and IV, defined above for the heat capacity isotherm and corresponding to the low-coverage two-dimensional condensation, may be absent, raising the possibility of a two-dimensional critical temperature. 4. Volume Bull and Breese (1968b) measured the specific volume of the ovalbumin-water system between 0.05 and 0.4 h. Above 0.2 h the partial specific volumes of water and protein are equal to those for the dilute solution. Below 0.2 h the partial specific volume of the water is 0.8 ml/g. Richards (1977) showed that, in partially dried crystals of human serum albumin, the partial specific volume of the solvent was constant and equal to the bulk value, to a hydration of 0.15-0.2 h (Fig. 5). Using dilatometry, Kim and Kauzmann (1980) measured the concentration dependence of the specific volumes of oxyhemoglobin, serum albumin, and ovalbumin at high concentrations (0.3-0.4 g/g of solution). Contributions on the order of ( c , ) ~ were not significant, indicating no effect of protein on solvent at concentrations at which the protein molecules are separated by a layer of water only 10-20 thick. This observation is consistent with a constant partial specific volume of the solvent over the concentration range studied. Dilatometric measurements of oxyhemoglobin and serum albumin are in accord with the above conclusion (Bernhardt and Pauly, 1975, 1980). 5. Ionization Infrared measurements by Careri et al. (1979b) show that the first water to bind to the dry protein leads to transfer of protons from carboxyl groups to other protein groups. Drying of the protein produces inversion of the pK order, and the effective pK of carboxylic acid side chains in the dry protein is above that of groups such as lysyl side chains, which,

51

PROTEIN HYDRATION AND FUNCTION

i

0

L

l

l

J

.2

.3

L

.4

I

.3

1

.6

.7

1

1

.8

.9

~

WEIGHT FRACTION of WATER

FIG. 5. Hydration dependence of the volume of human serum albumin crystals. Specific volume of crystals of the dimer form of human serum albumin as a function of water content. 0,Determined during drying; 0, determined during rehydration. Extrapolated = 0.734 cm3/g and = 0.995 cm3/g. The dashed line with intercepts of solid line i& arrowheads indicates the region of failure of the simple linear relationship and of deviation of partial specific volumes of protein and water from the dilute solution values. From Richards (1977).

in solution, are, of course, more basic. This can be understood as a way of reducing the electrostatic free energy associated with unsolvated charges. Apparently, the protein, when faced with a water-poor environment, abolishes as many charges as possible. The rate of hydrogen exchange in powders of lysozyme (Schinkel et al., 1985)is the same at 0.2 h as in dilute solution. Hydrogen exchange depends strongly on pH, and the effective pH in protein powders is controlled by ionization of the protein. Thus, at 0.2 h, corresponding to half-coverage of the surface, the effective pK of protein groups is that found in dilute solution.

52

JOHN A. RUPLEY AND GIORGIO CARER1

As noted above, normalization of the carboxylate ionization is likely the source of the reaction heat observed in region IV of the heat capacity isotherm. The carboxylate ionization process must contribute to the enthalpy isotherm in the low-hydration region. B . Denaturation The dependence of the denaturation process on hydration level has been studied by differential scanning calorimetry for several proteins, for example, P-lactoglobulin (Ruegg et al., 1975), lysozyme (Fujita and Noda, 1978, 1979), chymotrypsinogen A (Fujita and Noda, 1981a), and ovalbumin (Fujita and Noda, 1981b). Not surprisingly, in view of the long-known usefulness of dehydration for food preservation, the melting temperature (Td) increases sharply at low hydration (Fig. 6). The values of Td and AHd (Fig. 7) are equal or close to the solution values above 0.75 h, change slightly between 0.35 and 0.75 h (range II), and change strongly below 0.35 h (range I). Thermodynamic values estimated from scanning calorimetric experiments include a reaction-rate

1

I

60

70

80

r

90

1

I

100

TEMPERATURE [1oC]

FIG. 6. Thermograms of hydrated P-lactoglobulin from differential scanning calori.metry over the temperature range 60-105°C. Water contents (in grams per gram) and sample weights (in milligrams) are, respectively: (a) 0.176 and 6.275; (b) 0.439 and 8.562; and (c) 4.591 and 17.710. From Ruegg et al. (1975).

53

PROTEIN HYDRATION AND FUNCTION

1 " " " " " " l

390-

- 400

-0

E

LF

370-

2 -200 rc" a

350 -

l

0

n

~ 0.2

.

0.4

,

,

0.6

,

, , , 8 ~ 0.8 1.0 1.2

,

l

o

Water content (g g-1)

FIG.7. The temperature (Td)and enthalpy (Hd)of denaturation of ovalbumin as functions of the water content. From Fujita and Noda (1981b).

contribution, dependent on the scan rate, and the process studied may not be reversible. Results such as those shown in Fig. 7 have been interpreted as reflecting, in the higher hydration range 11, a secondary hydration phase, corresponding perhaps to the B shell of solvent about ions. In this view the water of range I1 would be a shell of solvent serving to interface the bulk solvent with the water ordered in the monolayer about the native protein surface. This molecular interpretation of the physics of the 0.35 to 0.75 h range conflicts with the heat capacity isotherms (see Section II,A,3), which show that the heat capacity of a native protein is invariant to hydration above about 0.4 h. The following alternative explanation of the scanning calorimetric measurements for the hydration range 0.35-0.75 h is more plausible, and possibly more interesting. The data of Fig. 7 describe the hydration dependence of the unfolding process, to which changes in hydration of unfolded species, as well as native species, contribute. If we understand the unfolded species to have a more extensive interface with solvent than the folded species, then the 0.75 h break point defines the point of full hydration of the unfolded species. This value is itself of interest and difficult to determine otherwise. Reducing hydration below 0.75 h necessarily destabilizes the unfolded relative to the native state, for which the hydration is constant above 0.4 h. The destabilization of the unfolded state results in a rise in T d .Presumably, the average conformation in the unfolded state changes as the hydration level is reduced. The unfolded state should be more compact and have more internal bonding at lower hydration levels, consistent with the decrease in AHd with decreasing hydration level.

54

JOHN A. RUPLEY AND GIORGIO CARER1

C. Nonfreezing Water The amount of water that does not freeze near 0°C in a protein-water system is a useful measure of the hydration water (Kuntz and Kauzmann, 1974). Such estimates are self-consistent, in that different methods for determining nonfreezing water give closely similar values, and are reliable, in that they agree with other thermodynamic estimates of hydration.

1 . Scanning Calorimetry Scanning calorimetric analysis of a sample containing enough water to freeze, when carried out over a temperature range including O"C, shows a large endotherm that is centered at or slightly below 0°C and corresponds to an ice-water phase transition. The amount of nonfreezing water associated with a weight of protein has been determined in two ways. Method I uses analysis of a set of samples of varied water content to define the largest amount of water that can be in the sample and yet not produce an ice-water transition endotherm. In Method I1 determination is made of the heat of the transition by integration over the endotherm and estimation of the amount of ice in the sample by use of the heat of fusion of bulk water, the nonfreezing water then being calculated by difference. Kuntz and Kauzmann (1974) reviewed early work (Berlin et al., 1970; Bull and Breese, 1968b; ,Haly and Snaith, 1971; Mrevlishvili and Privalov, 1969) on nonfreezing water in several systems, including the globular proteins serum albumin, ovalbumin, hemoglobin, and p-lactoglobulin. Ruegg et al. ( 1975) reinvestigated P-lactoglobulin, obtaining a threshold value of 0.29 h for the appearance of the ice endotherm. This value is more in agreement with the amount of nonfreezing water found for other globular proteins (0.3-0.35 h) than the value of 0.55 h reported by Berlin et al. (1970). Luescher et al. (1979) found 50 mol more nonfreezing water per mol of tosylated than native chymotrypsin. Ali and Bettelheim (1985)reported values of nonfreezing water two to three times greater than the values cited above and by Kuntz and Kauzmann (1974). 2 . Nuclear Magnetic Resonance (NMR) Kuntz, in a series of papers (see Kuntz and Kauzmann, 1974), developed the use of magnetic resonance as a probe of the freezing of solvent in protein solutions. The nonfreezing water in a sample measured at - 35°C appears as an NMR signal that is sharp compared to the broadband response for the ice phase. The NMR method gives results for proteins and other macromolecules in close agreement with estimates of

PROTEIN HYDRATION AND FUNCTION

55

nonfreezing water made by use of other methods, such as calorimetry (Kuntz and Kauzmann, 1974). Measurements of model polypeptides were consistent with the nonfreezing water being primarily associated with ionic groups of the protein (Kuntz, 1971). A set of amino acid hydration values, constructed to calculate the amount of nonfreezing water according to the amino acid composition of a protein, gave estimates in close agreement with measurement (Kuntz, 1971). Hsi and Bryant (1975) measured NMR relaxation for frozen lysozyme solutions. They found that the nonfreezing water consisted of two components with different T, values: The amount of the slower component was 0.06 g, and the amount of the faster component was 0.28 g of water per g of protein. The amount of the slower component, perhaps coincidentally, corresponds to the amount of tight-binding water seen in isotherm measurements. 3 . Diffraction and Other Methods

Poole et al. (1987) measured powder diffraction for lysozyme samples of varied hydration. Diffraction of ice crystallites was first detected at hydration levels greater than 0.3 h. The point of first appearance of frozen water is also defined by the shift and growth of the 0-D stretch band in infrared spectra measured for protein samples variously hydrated with deuterated solvent (Finney et al., 1982). Table I1 compares determinations of the nonfreezing water of lysozyme, measured by scanning calorimetry, NMR, infrared spectroscopy, and X-ray diffraction. TABLE II Corntarison of Nonfreezing Water, Determined by Several Methods"

Measurement

Nonfreezing water (grams of water per gram of protein) ~

Nuclear magnetic resonance

Differential scanning calorimetry Infrared spectroscopy Powder X-ray diffraction

~~

0.34b 0.34' 0.32 2 0.02d 0.32 2 0.02d 0.31 2 0.02d 20.30'

"All measurements shown are for hen egg white lysozyme. bFrom Kuntz (1971). CFromHsi and Bryant (1975). dData cited and analyzed by Finney et al. (1982). 'From Poole et al. (1987).

56

JOHN A. RUPLEY AND GIORGIO CARER1

4 . Comment

The determination of nonfreezing water is perhaps the most simple and straightforward way to estimate hydration. Scanning calorimetric and NMR measurements are made with equipment that is commonly available, and these methods should continue to be widely used. It is possible, however, that the nonfreezing water has no simple relationship to the interface water in protein powders or protein solutions. Nonfreezing water presumably results from competition between the protein surface and growing ice crystals for the water in the interface between the surface and the bulk solvent. Ice crystals would be expected to incorporate interface water over regions of the surface where water does not interact strongly with the protein or where the surface can accommodate the ice structure. Thus, the amount of nonfreezing water should be less than the hydration water, according to the extent of regions of the above type. The suggestion by Kuntz (1971), that the nonfreezing water is dominated by contributions from water about ionic residues, appears to conflict with the results of other measurements on partially hydrated proteins. These results suggest that ionic groups dominate as water sites in the tight-binding region of the isotherm, below the knee, and that polar protein groups dominate in the plateau region, which is of greater extent. D. Solutions 1. Hydration Forces

Two closely adjacent surfaces experience various forces: van der Waals, electrostatic, steric, and hydration. There are excellent review discussions of these interactions (Israelachvili, 1985, 1987; Israelachvili and Marra, 1986; Parsegian et al., 1985, 1986; Rand et al., 1985). Steric forces (Fig. 8B) arise from the thermal motions of surface groups, are statistical, and may have a large characteristic length, as is found for polymer chains bound to a surface. Hydration forces (Fig. 8A) arise from perturbarion of solvent by the surface; they may be propagate$ through many layers of water, with detectable interaction at 10-30 A distance. Using a surface forces apparatus, Israelachvili determined the force law for two molecularly smooth charged mica surfaces immersed in an aqueous solvent (see Israelachvili and Marra, 1986, and references cited therein). The repulsive hydration force is oscillatory (Fig. 8A). It is understood to reflect the geometry and local structure of the solvent and

PROTEIN HYDRATION AND FUNCTION

57 ENERGY (mJ/m2)

FORCE/RADIUS (mN/m)

'

DISTANCE (nm)

FIG.8. (A) Measured forces between two charged mica surfaces in M KCI, where beyond 30 8, (and out to 500 8,) the repulsion is well described by conventional electrostatic double-layer force theory. Below 30 8, there is an additional hydration repulsion, with oscillations superimposed below 15 A. (B) Forces between two uncharged lecithin bilayers in the fluid state in water. At long range there is an attractive van der Waals force, and at short range (i.e., below 25 8,) there is a monotonically repulsive steric hydration force. (C) Forces between two hydrophobized mica surfaces in water where the hydrophobic interaction is much stronger than could be expected from van der Waals forces alone. From lsraelachvili and Marra (1986).

to have the same origin as the radial distribution functions that describe pure liquids. Most simply, as two smooth surfaces are brought together in water, the natural geometry of the water is accommodated between the surfaces at some separations, but the fit is forced at other separations. This is consistent with the periodicity of the oscillation being 2.5 A. The oscillatory character is lost for rough surfaces or surfaces that undergo thermal fluctuations. Between surfaces separated by three layers of water-the two adjacent to each surface and one additional layer in between, for about 7.5 A separation of the surfaces-the hydration force calculated for the water of the in-between layer is 10- 100 cal/mol. The hydration force decays exponentially with increase in separation. Parsegian, Rand, and colleagues (Parsegian et al., 1986; Rand et al., 1985) developed an elegant and simple method of determining the hydration force as a function of distance between surfaces, based on the observation that the hydration force and the surface separation each depend on water activity, and these dependences can be measured. A sample containing aggregates with extensive surfaces (e.g., lipid bilayers or DNA) is placed in an environment where water activity can be varied. As the water content of the sample changes according to the water activity, the volume and thus the spacing between the surfaces change.

58

JOHN A. RUPLEY AND GIORGIO CARER1

The spacing is measured in separate experiments, as by use of X-ray diffraction. Furthermore, the force corresponding to the spacing is proportional to the osmotic pressure, which is, of course, also a function of the water activity. The water activity can be controlled conveniently by establishing osmotic equilibrium with a large volume of a solution of a polymer (e.g., dextran) at an appropriate concentration. This allows accurate specification of small osmotic pressures for water activities near unity, corresponding to the relatively small energies per water molecule associated with hydration forces. Figure 9 shows force laws for phosphatidylcholine bilayers (Lis et al., 1982), determined by the osmotic stress method. Similar data were obtained for DNA samples (Rau et al., 1984). The characteristic length governing decay of the force is about 3 A for both systems. Interactions of this kind can also be important for protein aggregates. Prouty et al. (1985) used the osmotic stress method to determine the phase diagram of sickle cell hemoglobin (Fig. 10). At a critical osmotic pressure, which is temperature dependent, a solution of deoxyhemoglobin S collapses to a gel, with a large change in volume. One of the strengths of the osmotic stress method is that it provides additional information that can be used for thermodynamic analysis of the system.

1000 100 10 1

.l .01

0

5

10

IS

20

Slayer Separation

25

30

(A)

FIG.9. Force-distance relationshipsfor lipid bilayers. Data for repulsion between dilauroylphosphatidylcholine bilayers at 25°C. At high pressures ( 0 )the bilayers have been forced into a frozen-chain gel phase, a response that shows the structural importance of forces exerted by osmotic stress. The exponential part of the melted liquid-crystalline samples ($7) is best fit by an exponential decay constant of 2.6 A. From Parsegian el al. (1986).

59

PROTEIN HYDRATION AND FUNCTION

4c

3c

0,

I

-E

2c

*’

10

I

C

100

I

I

1

I

I

200

300

400

500

600

Molor volume ( 1

FIG. 10. Osmotic pressure-molar volume isothermals for deoxygenated sickle hemoglobin (Hb). (Inset) Pressure-volume isotherms for CO,, taken from a standard physical chemistry text, show the resemblance to classical gas condensation. From Prouty et al.

(1985).

Claesson et al. (1989) measured the forces between hydrophobized mica surfaces, with and without adsorbed insulin. They concluded that the range of the hydration force for this globular protein is less than 10 A and that the hydration layer is not more than one or two water molecules thick. Hydration and other surface forces are important for the approximation of aggregates with large areas. It is possible that the concept of

60

JOHN A. RUPLEY AND GIORGIO CARER1

hydration force, defined in terms of interaction between surfaces, may not have meaning for the hydration of an isolated protein molecule or for the interaction of a protein with a small molecule. Specifically, hydration forces reflect the special geometry of very large smooth surfaces being brought close together, and an isolated surface might produce no significant long-range perturbation of the surrounding solvent. Even if one allows that hydration forces are of undiluted importance for an isolated surface, the perturbation of the chemical potential of the water in the second layer is an order of magnitude less than that in the first layer, and the perturbation decays exponentially with distance from the surface.

2. Preferential Solvation and Multicomponent System There is a substantial literature on the thermodynamics of threecomponent systems-water, protein, and second solute. For a review of early work, methods, and theory, with emphasis on sedimentation experiments, see Kuntz and Kauzmann (1974). Timasheff and colleagues (see Lee et al., 1979, and references cited therein) have developed a beautiful formalism for treating the thermodynamic nonideality of threecomponent systems in terms of the preferential interaction parameter

where g, is the weight concentration of protein (component 2) or second solute (component 3). A negative value for the preferential interactionnot uncommonly found for salts, sugars, and polyols-corresponds to the region about the protein being depleted in solute relative to the bulk solvent, that is, to preferential hydration. As a rule solutes that stabilize the native conformation show a negative value for the preferential interaction parameter, probably owing to the greater hydration and an even more negative interaction parameter for the denatured protein. Winzor and Wills (1986) have shown that the analysis of nonideality in terms of preferential solvation is equivalent to an alternative analysis based on excluded volume. T h e excluded volume model is commonly applied to the nonideality of solutions of macromolecules, and it is rooted in the statistical mechanics of polymer solutions, equations of state, and virial expansions. Various other physical methods have been applied to the study of protein hydration in multicomponent systems, for example, NMR spectroscopy of frozen samples (Izumi et al., 1980), scanning calorimetry (Fujita et al., 1982), thermodynamics of denaturation (Velicelebi and Sturtevant, 1979), and sorption (Stonehouse, 1982). A protein crystal is a well-defined multicomponent system, which is

PROTEIN HYDRATION AND FUNCTION

61

generally well behaved, in that the density is linear in composition of the mother liquor (Matthews, 1985). Early work aimed at defining the amount of hydration water used this observation (Adair and Adair, 1936; Perutz, 1946). Scanlon and Eisenberg (1975, 1981) refined the thermodynamic analysis and estimated the hydration for myoglobin and several other proteins from the dependence of the crystal density on the density of the mother liquor, with the additional assumption that the solvent not part of the hydration shell had the density of the mother liquor. Values estimated for the hydration ranged from 0.05 to 0.27 h, and values for the specific volume of the hydration water ranged from 0.9 to 1.9 ml/g. The model used in this treatment appears to be similar to the model used for analysis of preferential hydration, and in both cases the values estimated for the hydration water vary substantially between proteins.

3 . Compressibility Kundrot and Richards (1987) compared crystal structures of hen egg white lysozyme determined at 1 and 1000 atm. The crystallographically determined water sites were similar in the two structures. T h e same authors (Kundrot and Richards, 1988) reported the dependence of the density of the crystals on hydrostatic pressure, which, with the crystallographic information, gave an estimate of the compressibility of the crystal water. This was equal, within the experimental error of about 1576, to the compressibility of bulk solvent. Considering the uncertainty, the compressibility of the hydration water is in accord with measurements of the heat capacity, volume, and enthalpy, that showed 10- 15%differences between interface and bulk solvent. The compressibility of the protein, estimated from the crystallographic data, was about one-half of the adiabatic compressibility determined in solution measurements (see Gavish et al., 1983, and references cited therein). T h e p sheet domain was less compressible than the other lysozyme domain, within which the structural elements (e.g., the helices) responded complexly to change in hydrostatic pressure. Kundrot and Richards (1988) and Gavish et al. ( 1983) discussed the implications of protein compressibility values for protein fluctuations. 111. DYNAMICS A . Dielectric Relaxation

The strongly polar water molecule moves in response to an alternating electric field. This process, dielectric relaxation, is a useful probe of the

62

JOHN A. RUPLEY AND GIORGIO CARER1

properties of water in biological materials. The dielectric and electronic properties of proteins have been reviewed by Pethig (1979; Pethig and Kell, 1987) and, for the microwave region, by Parak (1986). The investigation of protein hydration in solution by dielectric methods has been described by Fel'dman et al. (1986) and by Grant et al. (1985). The latter reviews progress made in the field since a previous discussion of the possibilities and limitations of the method (Grant, 1982). Kuntz and Kauzmann (1974) reviewed early work on dielectric relaxation in protein solutions and powders, as it bears on protein hydration. The measurements described in this section are for powder samples, unless otherwise indicated. 1 . High Frequency

Harvey and Hoekstra (1972) determined the dielectric constant and loss for lysozyme powders as a function of hydration level in the frequency range 10'- 1O1OHz. At water contents less than 0.3 h, they found a dispersion at 170 MHz, which increased somewhat with increasing hydration, and a new dispersion at about 1O1O Hz that develops at high hydration. These dispersions, detected by time-domain techniques, remain measurable down to the lowest temperature studied, - 60°C. Water mobility in the hydration shell below 0°C is in line with other observations of nonfreezing water. Above 0.3 h, in the stage of the hydration process at which condensation completes the surface monolayer, water motion increased strongly with increased hydration (Fig. 11). Singh et al. (1981) measured the relaxation time for the water of a sample of myoglobin crystals in the microwave region over a wide range of temperature. No discontinuity in the dielectric relaxation rate was observed near 273 K, indicating that essentially no free water was present in the crystals, that is, nearly 400 molecules of water per molecule of protein were perturbed. The dielectric relaxation rate at 200 K was two orders of magnitude less than at room temperature (Fig. 12). The data correlate well with the motional properties of the heme iron measured by Mossbauer spectroscopy. Singh et al. (1981) suggested that there is electrical coupling of protein and solvent motions. Kent and Meyer (1984) made broad-band dielectric measurements in the frequency range 0.3- 16 GHz on various hydrated protein powders, including hemoglobin and bovine serum albumin. Comparison of relaxation spectra measured at 20-80°C suggested that at high hydration there is more than one state of multilayer water. Genzel et al. (1983) and Kremer et al. (1984) reported picosecond relaxations in proteins, including lyophilized hemoglobin and lysozyme, that were described in terms of processes occurring in asymmetric double-well potentials, likely the NH OC hydrogen bridges of the

...

1

.'

- - FIG. 1 1 . Hydration dependence of dielectric response at 25 GHz. Dielectric constant ( E ' ) and loss ( E " ) of packed lysozyme powder as a function of water content. Frequency, 25 GHz: temperature, 25°C. (From Harvey and Hoekstra, 1972.)

FIG. 12. Low-temperature dielectric response at 10 GHz. Dielectric relaxation rates of water in metmyoglobin (O),free water (0), and ice (A). From Singh et al. (1981).

64

JOHN A. RUPLEY AND GIORGIO CARER1

peptide backbone. Poglitsch et al. ( 1984) measured dielectric absorption, in the region 50-150 GHz, for lysozyme at different hydration levels. No additional absorption due to hydration water was detected. The frequency range of these measurements is higher than that covered by Singh et al. (198l),who detected hydration water. Shchegoleva (1984) investigated the hydration dependence of the dielectric constant at 7.6 mm wavelength for various other globular proteins and biopolymers.

2. KHz and M H z Frequencies Hawkes and Pethig (1988) identified a weak dielectric loss in the KHz region and explained it in terms of vibrational motions of the polypeptide backbone. Using time-domain reflectometry, Bone (1987) observed for chymotrypsin a dielectric dispersion that was centered at 12 MHz and increased with increasing hydration. The dispersion was attributed to relaxation of polar elements of the protein. Careri et al. (1985) measured dielectric losses for lysozyme powders at varied hydration level in the 10-KHz to 10-MHz frequency range, by use of a dielectric-gravimetric technique. The isotope effect and pH dependence indicated that the conduction process was protonic. The binding of an oligosaccharide substrate [(GlcNAc),] increased the characteristic time for the relaxation, to an extent equivalent to a 2-fold reduction in the inferred d.c. conductivity (Fig. 13). These observations lead to two conclusions. (1) Half of the proton flow of the conduction process passes through the active site, which comprises only one-tenth of the protein surface; apparently, the active site is special in facilitating the movement of protons. (2) The conduction process is cooperative, judged by the seventh-order dependence on hydrogen ions bound. Careri et al. (1985) used a three-layer composite capacitor, made of glass, protein sample, and air, without direct contact between electrodes and sample. The frequency-dependent relaxation is due to MaxwellWagner polarization (Pethig, 1979; Pethig and Kell, 1987), which is the interfacial polarization displayed by a heterogeneous medium and is due to the different d.c. conductivities and d.c. dielectric constants of the constituent components of the medium. It is coincidental that the effects measured by Careri et al. (1985) fall in the frequency range of the data of Hawkes and Pethig (1988) and Bone (1987), who measured intrinsic frequency-dependent dielectric properties of the protein sample, for a capacitor consisting of only the protein sample. Careri et al. (1986), using the framework of percolation theory, analyzed the explosive growth of the capacitance with increasing hydration above a critical water content (Fig. 14). The threshold for onset of the dielectric response was found to be 0.15 h for free lysozyme and 0.23 h for the lysozyme-substrate complex. In the percolation model the thresh-

h

.,

FIG. 13. Hydration dependence of protonic conduction. The dielectric relaxation time, 76, is shown versus hydration, h, for lysozyme powders. The relaxation time is proportional to the reciprocal of the conductivity. (A) H,O-hydrated samples: solid curve, lysozyme without substrate; lysozyme with equimolar (GICNAC)~ at pH 7.0; 0, with 3 X molar (GlcNAc)l at pH 6.5. The relaxation time is nearly constant between pH 5.0 and 7.0. (B) *H*O-hydrated samples: solid curve, lysozyme without substrate; 0 , lysozyme with equimolar (GIcNAc)~ at pH 7.0. From Careri et al. (1985).

8.0

0.0010 -E

7.0 -

10 kHz

400 kHz

6.0 5.0 -

G P

0

-

4.0

3.0 2.0

1 .o

C ._

E - 0.0005 9

1

I

0.0

2 MHz

.+;*.*;::;*;

.

,

I;**::

0.3 0.4 h (g of H20 per g of dry weight)

0.1

0.2

.

UI

1 0.0000

0.5

FIG. 14. Hydration dependence of capacitance [O; C, in picofarads (pF)] of the composite capacitor containing a sample of lysozyrne powder of pH 3.1 1 as a function of hydration level of the protein. The capacitance data are given for three frequencies. The hydration level was decreased from the high-hydration limit of more than 0.35 h to the low-hydration limit of near 0.07 h by passage of a stream of dry air through the apparatus. The evaporation rate E (0;grams of water evaporated per minute) decreases to 0 at the low-hydration limit. From Careri et al. (1986).

66

JOHN A. RUPLEY AND GIORGIO CARER1

old corresponds to the point of appearance of a cluster of conducting elements, called the infinite, or unbounded, cluster, that connects all regions of the system. Immediately above the threshold the probability that a conducting element is part of the infinite cluster increases strongly, leading to much richer interconnections within the infinite cluster, and so to a correspondingly strong increase in the conductivity, as observed. The fractional coverage of the surface at the onset of protonic conduction is 0.40, which is in close agreement with the theory for surface percolation that predicts a value of 0.45. The critical exponent describing the growth of the conduction process above the critical point is t = 1.29 (Careri et al., 1988), also in close agreement with the theory for surface percolation (Clerc et al., 1980; Stauffer, 1985; Zallen, 1983). For the lysozyme-saccharide complex the threshold for enzyme function (Rupley et al., 1980) is the same as the threshold for establishing percolation and long-range connectivity. Although binding of substrate shifts the percolation threshold from 0.15 to 0.23 h, the critical exponent is unchanged. This approach has been extended by Rupley et al. (1988) to study of the water-induced percolation in hydrated purple membrane fragments of Halobacterium halobium. The results and conclusions are qualitatively similar to those reported above for lysozyme. (1) The percolation is twodimensional, judged by the value of the critical exponent (Fig. 15). (2) Certain regions of the surface provide preferred protonic conduction paths. (3) There is a correspondence between the onset of function-here, the photoresponse-and the establishment of long-range connectivity within the surface water clusters. Table I11 summarizes results for the critical exponents and critical coverages, obtained by percolation theory analysis of the protonic conduction processes for lysozyme and purple membrane, and compares these values with theory. 3 . Low Frequency

Eden et al. (1980), Gascoyne et al. (1981), and Bone et al. (1981) measured the hydration dependence of the d.c. conductivity and dielectric properties at frequencies to 33 GHz for compressed samples of bovine serum albumin and lysozyme. The data for low frequency were interpreted in terms of activated hopping of a fixed number of charge carriers, which were considered to be protons originating from ionizable carboxylic acid groups and moving along a network of water-protein and water-water bonds. Further work by Bone and Pethig (1982, 1985) and Behi et al. (1982) confirmed this conclusion and extended it to other proteins. The low-frequency dielectric properties vary with the pH and the hydration level of the powder sample. Direct measurements of solid-

67

PROTEIN HYDRATION AND FUNCTION

-3.5

-3.0

-2.5

-

-2

.o

-1.5

Log ( h - h c )

FIG. 15. Critical exponent for protonic percolation on purple membrane. Hydration dependence of the conductivity for H 2 0 (0)and * H 2 0 (0)hydration of lyophilized samples of fragments of purple membrane from Halobacterium hulobium. Only data near the percolation transition, with values of (h - h,) < 0.01, were plotted. The critical exponent, t, determined from the slope of the lines, is 1.23 for both hydration regimens. The ratio of the rates in normal and deuterated water is 1.38, in close agreement with the square root of the mass ratio. From Rupley et al. (1988). TABLE 111 Percolation Parameters for Lysozyme, Purple Membrane, and Other System a System Lysozyme Hydrated with H 2 0 Hydrated with D 2 0 1 : 1 Complex with (GIcNAc)~, hydrated with H 2 0 Purple membrane Hydrated with H 2 0 Hydrated with D 2 0 Theory Two-dimensional percolation Three-dimensional percolation

Fractional Critical Coverage

Critical Exponent

0.37 0.41

f

f

0.04 0.01

1.30 & 0.09 1.24 f 0.08

0.58

f

0.01

1.34 f 0.11

0.18 0.18

1.231 1.232

0.45 0.15

1.1-1.3 2.0

“Data for lysozyme are from Careri et al. (1988). Data for purple membrane are from Rupley et al. (1988). For summaries of theoretical analyses see Stauffer (1985), Zallen (1983), and Clerc et al. (1980). The critical exponent can be considered a truer test of the order of the percolation process than the fractional critical coverage.

68

JOHN A. RUPLEY AND GIORGIO CARER1

state conduction and other dielectric results have been reported by Morgan and Pethig (1986), who concluded that the low-frequency dielectric dispersion had to be associated with interactions between ions and metal electrodes. Ataka and Tanaka (1980) measured the d.c. conductivity of lysozyme single crystals at different temperatures. They attributed their results to protons originating from residual water molecules. The contact between electrode and crystal was made with silver paste, a possible flaw in the experimental method. Shablakh et al. (1984) investigated the dielectric properties of bovine serum albumin and lysozyme at different hydration levels, at low frequency. Besides a relaxation attributed to the electrode-sample interface, they detected a further bulk relaxation that can be confused with a d.c. conduction effect. The latter relaxation was explained by a model of nonconductive long-range charge displacement within a partially connected water structure adsorbed on the protein surface. This model has nonconventional features that differ from the assumptions of other more widely accepted models based on Debye relaxations. Tredgold et al. (1976) measured the polarization of hydrated films of lysozyme at 0.1 Hz and concluded that the apparent high dielectric constant was attributable to protonic conduction in the water of crystallization. 4. Thermal Depolarization Thermally stimulated depolarization currents are detected in a sample first cooled to low temperature in a capacitor with shorted electrodes, then warmed slowly with the electrodes connected to a sensitive d.c. electrometer. In this way the dielectric relaxation processes occurring in the sample are displayed separately, according to their activation energies and barrier heights, during the scan over temperature. Celaschi and Mascarenhas (1977) studied nearly dry lysozyme by electret thermal depolarization, thermal-stimulated pressure, isothermal polarization decay, and thermogravimetry. For a change in temperature of the sample from 250 K to room temperature, desorption of water dipoles was the main process responsible for electrical depolarization. Depolarization thermal studies have been reported by Leveque et al. (1981) for partially hydrated keratin, by Bridelli et al. (1985) for lysozyme, and by Anagnostopoulou-Konsta and Pissis ( 1987) for casein. These studies reveal a rich and complex thermal depolarization spectrum, shown in Fig. 16. It is difficult to explain the spectrum with a model based on the reorientation of noninteracting dipoles with a few distinct relaxation times. Pissis and colleagues (personal communication

PROTEIN HYDRATION AND FUNCTION

69

30-

z- 20

-

10

-

‘T

T

IKI

FIG. 16. T h e high-temperature thermally stimulated depolarization current band for casein samples with eight different water contents: A, h = 0.013; B, h = 0.04; C, h = 0.067; D, h = 0.09; E, h = 0.10; F, h = 0.124; G , h = 0.164; H , h = 0.216. From Anagnostopoulou-Konsta and Pissis (1987).

to G. Careri, 1988), working with hydrated lysozyme, have detected a thermally stimulated depolarization band that grows with increasing hydration and is associated with the same threshold hydration level as the one detected for protonic percolation by Careri et al. (1986).

B . Percolation Model The use of the percolation model to analyze the d.c. conductivity in hydrated lysozyme powders (Careri et al., 1986, 1988) and in purple membrane (Rupley et al., 1988) introduces a viewpoint from statistical physics that is relevant to a wide range of problems originating in disordered systems. Percolation theory is described in the appendix to this article, for readers unfamiliar with it. Here, we discuss the significance of percolation specifically for protein hydration and function. One can picture the percolation process detected in the dielectric response as proton transfer along a thread of hydrogen-bonded water molecules adsorbed on the protein surface (Careri et al., 1986). The water molecules are formally equivalent to the conducting elements of the familiar percolation model of a conducting network. Above the thresh-

70

JOHN A. RUPLEY AND GIORGIO CARER1

old the long statistical path of interconnected water molecules acts as a “short” that bypasses the local geographical details, through displacement of protons the length of the macromolecule. The statistical character of the percolation model deserves emphasis. The model is based on the random arrangement of elements over a surface. For an ensemble of surfaces, or for the time evolution of a single surface, one can view the clusters and thus the paths of connected elements as fluctuating. Percolation through the protein interior appears to be unlikely; the internal water in proteins is sparse and unable to contribute to longrange connectivity. Protonic percolation is likely to be similar for all globular proteins for the following reasons: percolation is insensitive to local details of structure; the sorption isotherms of globular proteins are nearly identical; and the essential features of a percolation transition are size independent. The thermodynamics of the hydration process define two phase transitions (see the discussion in Section I1 of the heat capacity and other thermodynamic properties). The percolation transition is fundamentally different from these transitions. The former, and most other familiar phase transitions, are associated with a discontinuity in structure and chemistry of the system. The percolation transition, in contrast, is detected as a change in a process, specifically, a long path length proton movement. At the critical point for percolation, there is a discontinuity in the connectivity, but no discontinuity in the structure or chemistry of the system. In a pure percolative system there is no energy of interaction between elements, and the essential feature of the transition is the establishment of the infinite (unbounded) cluster, connecting all regions of the system. For a protein the percolative transition falls in a region where there is no discontinuity in the thermal properties (i.e., heat capacity and enthalpy). As expected, the energy of interaction between protein surface elements is not an essential feature of protonic percolation. Since a change in surface coverage of about 2% (6 mol of water per mol of lysozyme) shifts the system from the nonconducting to conducting mode, one can envision biochemical control based on percolation. Conduction in membranes might be turned on or off by adding or subtracting a few water molecules or other conducting elements, without a need for change in protein or membrane conformation. As noted, for both the lysozyme-saccharide complex and the purple membrane, the critical point for protonic percolation is at the onset of function. These observations may apply to other situations, in which a new property emerges suddenly at a critical water content, and may lead to understanding of function in terms of the building up of a statistical network of water-assisted pathways encompassing the system. Statistical

PROTEIN HYDRATION AND FUNCTION

71

long-range connectivity should be considered along with other intrinsic properties of proteins (e.g., charge distribution, pH, and conformation at the active site) when analyzing function within the framework of statistical physics. In this context measurements of protonic conduction can be viewed as an experimentally convenient probe of the existence and properties of extended hydrogen-bonded networks on the protein surface.

C . Resonance 1 . Nuclear Magnetic Resonance This section discusses a selection of NMR results with an emphasis on powder studies, on experiments that describe the dynamics of water at the protein surface, and on lysozyme as a model protein. Methods and theory are not discussed. For review discussions see Kuntz and Kauzmann (1974), Bryant (1978), Koenig (1980), and Fung (1986). A recent review by Bryant (1988) is an elegant summary of the theory and results for NMR measurements of protein hydration, in powders and in solution. a. Powders. Bryant and collaborators have carried out an extensive series of measurements on powder samples of lysozyme. Hilton et al. (1977) examined the relaxation of water as a function of temperature in powders of varied hydration level. The experiments paralleled the dielectric dispersion measurements of Harvey and Hoekstra ( 1972).Water near the protein surface responded as a fluid, with motion decreasing as the hydration level decreased (Fig. 17). The reorientational correlation time was reduced by less than a factor of 100 from the value for bulk water (Bryant and Shirley, 1980a,b). The data are consistent with a model in which the surface water is localized, in which water rotation is about the protein-water bond, and in which there is motional averaging to include also a slower reorientation of the rapid-motion axis (Shirley and Bryant, 1982). Peemoeller et al. (1981, 1984, 1986), using a twodimensional method, carried out *H magnetic resonance studies, for which water-protein cross-relaxation is absent. At half-saturation of the lysozyme surface with water, about 90% of the adsorbed water has the axis of fast rotation colinear with the water-protein hydrogen bond. The correlation time for rotation about this axis is 10-9-10-’0 sec (1/100th of the bulk water value), and that for reorientation of the axis is lO-’sec. Andrew and collaborators (Andrew, 1985; Andrew et al., 1983; Gaspar et al., 1982) measured the temperature and hydration dependence

72

JOHN A. RUPLEY AND GIORGIO CARER1

2ok 00

10

g Hfl/100 g PROTEIN

FIG. 17. Longitudinal 'H NMR relaxation parameters at 30 MHz for water adsorbed on lysozyme powders derived from the cross-relaxation model after setting the protein relaxation rate equal to 0. T I , is the water proton relaxation time and T , is the time constant characterizing spin transfer between the protein protons and the water protons. From Hilton et al. (1977).

of relaxation for several macromolecules. Results for lysozyme are given in Fig. 18. The most strongly adsorbed water, below the knee of the isotherm, is associated with a large temperature-dependent NMR response to change in hydration level. The commonly observed transition in dynamic behavior near 200 K is displayed in the NMR data. b. Solutions. 'H, *H, and I7O magnetic relaxation measurements on aqueous solutions of lysozyme and several other proteins, made as a function of frequency, provide information about the dynamics of surface water in the fully hydrated state (Bryant et d.,1982; Halle et d., 1981; Koenig, 1980; Koenig and Schillinger, 1969; Koenig et al., 1975; Piculell and Halle, 1986). I7Orelaxation is a particularly powerful probe, as it lacks the complications of cross-relaxation between protein and solvent protons and of exchange of labile hydrogens of the protein with solvent. Comparisons of I7O and 2H data show that exchange of labile hydrogens makes a significant contribution. The picture of surface dynamics that emerges (Halle et al., 1981; Koenig, 1980) is similar to that described above for partially hydrated solid protein samples: fast rotational motion for the solvent, perhaps 10 times below the bulk solvent rate; a slower reorientation time of about 10 nsec; and few, if any, waters that are immobile on the NMR time scale. More water enters into the NMR response than detected by perturbation of thermodynamic properties.

PROTEIN HYDRATION AND FUNCTION

73

FIG. 18. Temperature and hydration dependence of NMR relaxation. Variation with temperature of the proton spin-lattice relaxation time, T ,, at 60 MHz of polycrystalline lysozyme with various degrees of hydration. H, hydration with H 2 0 ; 0---0, hydration with D20.From Andrew (1985).

By measurement of nuclear Overhauser effects for pancreatic trypsin inhibitor in solution, Otting and Wuethrich (1989) identified the four structural water molecules found in the crystal by diffraction measurements. Other hydration waters exchanged rapidly with bulk solvent, that is, with a proton-exchange lifetime shorter than 3 x 10-*0sec. Polnaszek and Bryant (1984a,b) measured the frequency dependence of water proton relaxation for solutions of bovine serum albumin reacted with a nitroxide spin label (4.6 mol of nitroxide per mol of protein). The relaxation is dominated by interaction between water and the paramagnetic spin label. The data were best fit with a translational diffusion model, with the diffusion constant for the surface water in the immediate vicinity of the nitroxide being five times smaller than that for

74

JOHN A. RUPLEY AND GIORGIO CARER1

bulk water. A slightly lower diffusion constant was estimated by Schauer et al. (1988), from an analysis of 2H relaxation data, by use of a model that included translational diffusion. Shimanovskii et al. ( 1977) used paramagnetic interaction to characterize other aspects of the surface water. Bryant and collaborators (Borah and Bryant, 1982; Hsi and Bryant, 1977; Hsi et al., 1976a,b) carried out 'H or 2H magnetic resonance measurements on crystals of lysozyme and chymotrypsin. Usha and Wittebort (1989) studied the 2H NMR of crystalline crambin. At 140 K the protein hydrate is stationary, with T = 1 msec. Above 200 K changes in the signal with temperature are consistent with a glass transition or melting of the hydration water. This broad transition parallels closely the changes with temperature found for the heat capacity, Mossbauer spectroscopic, and other properties of hydrated protein crystals. At room temperature no more than 12 water molecules are orientationally ordered. The average rotational correlation time of the hydration water is about 40 times longer than that for bulk water. c. Amount of Hydration Water. Pessen and Kumosinski (1985) derived expressions for the protein concentration dependence of the 2H and 'H relaxations of protein-salt solutions. Their estimates of the amount of slow-tumbling water associated with (P-lactoglobulin ranged from 0.005 to 0.04 g of water per g of protein, under a two-state fast-exchange model and a three-state model. Bourret and Parello (1984) estimated, from 'H NMR measurements, that 110 water molecules are bound to polar surface sites of lysozyme. Sloan et al. ( 1973) titrated glyceraldehyde-3-phosphate dehydrogenase with nicotinamide adenine dinucleotide (NAD) and observed an increase in water relaxation by about 25% over that from the protein alone. They interpreted this effect as an increase of at least 26 mol of hydration water per mol of protein. This conclusion contrasts with a volume contraction and decrease in preferential hydration observed through other measurements to be associated with binding of NAD (Durchschlag et al., 1971; Sloan and Velick, 1973). Fullerton et al. (1986) measured 'H spin-lattice relaxation during dehydration of lysozyme solutions to a nearly dry state, and during rehydration of lyophilized lysozyme powder by isopiestic equilibration and, for high hydration levels, by titration with water. Breaks in the NMR response were found at 0.055, 0.22-0.27, and 1.22-1.62 h (Fig. 19 shows the two higher hydration discontinuities in slope). Estimates of the water correlation times are 2 x and 5 X lo-" sec, respectively, for the three classes of water defined by the breaks. The 0.055

75

PROTEIN HYDRATION AND FUNCTION

22-

x x x

x o

x

20-

18-

16l4-

l2-

5

lo86-

0

1

2

3

4

5

6

7

8

9

0

Ms/h.l FIG. 19. A dehydration from dilute solution study of the spin-lattice relaxation rate ( l / T l )versus concentration [mass solute/mass water (Ms/M) = h-’1 for six different initial concentrations of lysozyme: 0, 0.5 g/ml; A, 1.0 g/ml; 0, 1.5 g/ml; 0 , 2.0 glml; 0 , 2.5 glml; and x , 5.0 g/ml. The lack of dependence on initial concentration shows that “equilibration”time is not an important parameter. From Fullerton et al. (1986).

and 0.22 h discontinuities in slope are at the hydration levels of discontinuities in various other dynamic and time-average properties. Lioutas et al. (1986) measured the I7Oand *H resonances of lysozyme powders and solutions, in experiments like those carried out for ‘H by Fullerton et al. (1986). They similarly interpreted discontinuities in the NMR response in terms of three populations of water: 20 rnol of water per rnol of protein (corresponding to 0.025 h) with a correlation time of 41 psec, 140 mol of water (0.17 h) with a correlation time 27 psec, and 1400 mol of water (1.7 h) with a correlation time 17 psec. T h e differences between these results and those of Fullerton et al. (1986) indicate the difficulty of estimating water correlation times. Lioutas et al. (1987) extended these results by analyzing ’H resonance data through comparison with the sorption isotherm. D’Arcy-Watt analysis of the sorption isotherm gave 19 mol of tightly bound water per mol of lysozyme, 148 mol of weakly bound water, and 2000 mol of multilayer water. These classes plus two more types, corresponding to water in solutions

76

JOHN A. RUPLEY AND GIORGIO CARER1

of lysozyme dimer and monomer, respectively, were sufficient for explaining the hydration dependence of the resonance signals. Halle et al. (1981) measured 1 7 0 NMR relaxation for solutions of several proteins as a function of frequency and protein concentration. They estimated hydration by use of a two-state fast-exchange model with local anisotropy and with assumed values of the order parameter and several other variables. The hydration values ranged from 0.43 to 0.98 h for five proteins, corresponding approximately to a double layer of water about a protein. The correlation time for water reorientation was, averaged over the set of proteins, 20 psec, about eight times slower than that for bulk water. A slow correlation time of about 10 nsec was attributed to an ordering of water by protein at very high concentration. Kakalis and Baianu (1988) obtained similar results for lysozyme. They estimated 180 mol of hydration water per mol of lysozyme in the absence of salt. In 0.1 M NaCl solution the hydration was 290 mol/mol. If one were to draw a consensus from the experiments described above, it would be that perhaps two layers of water about the protein, corresponding to about 1 g / g of protein, are affected in NMR behavior by the protein and that this water has perhaps 10 times slower motion than bulk water. T h e NMR relaxation behavior of a protein-water system is, however, complex, particularly with regard to 'H relaxation. Koenig (1980) and Eisenstadt (1985) gave excellent discussions of the difficulties and of what can be safely concluded. The extraction of an estimate of hydration from NMR measurements requires assumption of a mechanism for the relaxation process, and generally also specification of some parameter values based on other information. In view of the uncertainty attached to the models, it is difficult to obtain acceptable quantitative estimates of hydration. It is certain that there are few, if any, waters with long residence times [e.g., hemoglobin has fewer than five waters with correlation time longer than sec (Eisenstadt, 1985)]. 2 . Electron Spin Resonance (ESR) Likhtenshtein and colleagues (Belonogova et al., 1978, 1979; Likhtenshtein, 1976) carried out a series of measurements on the hydration dependence of the mobility of spin labels covalently bound to several proteins. T h e results were correlated with Mossbauer spectroscopic data obtained in parallel experiments. Spin-labeled human serum albumin and a-chymotrypsin showed a critical hydration level for onset of motion at relative humidity 0.8, equivalent to 0.2 h. The temperature dependence of the spin label spectrum showed a critical temperature of 230 K, below which motion was frozen. Serum albumin labeled at surface sites

PROTEIN HYDRATION AND FUNCTION

77

with 57Fegave Mossbauer spectra with a different hydration dependence (critical hydration level at 0.6 relative humidity), and with a different temperature dependence (motion frozen at 200 K). Chymotrypsin with surface Mossbauer labels showed a critical hydration level and temperature like those found for the spin-labeled protein. Hemoglobin and ferredoxin showed a critical hydration level for spin label mobility at 0.6 relative humidity, and for Mossbauer mobility at 0.5 relative humidity. For the latter two proteins the 57Felabels were incorporated into the heme of hemoglobin and the iron-sulfur cluster of ferredoxin, and were buried within the protein matrix. There appears to be generally good, but not exact, agreement between the hydration dependences of the Mossbauer and spin labels. That they are so similar is perhaps surprising, considering the different motions sampled by the two techniques. The rotational correlation times of the spin labels range from sec for the dry or low-temperature samples to lo-* sec for the wet and room temperature samples. The later characteristic time is an order of magnitude or more below that for water motion in comparably hydrated samples (see above) or for motion of a noncovalently bound spin probe (see below). The rotational motion of a spin label is determined in part by the dynamics of the site of attachment. Steinhoff et al. (1989) measured the temperature and hydration dependence of the ESR spectra of hemoglobin spin-labeled at cysteine p-93. They observed the critical temperature near 200 K, as described above, and the sensitivity of the spectrum to hydration level. Spectrum simulations suggested that there were two types of motion: in the dry protein, a fast vibration of the label within a limited motion cone; upon the addition of water, a hydration-dependent motion assigned to the fluctuations of the protein, of correlation time sec in samples of high hydration and at 300 K. The temperature dependence of the motional properties of a spin probe (TEMPONE), diffused into hydrated single crystals, closely paralleled the motional properties of the label. This was taken to be evidence for coupling between the dynamical properties of the protein and the adjacent solvent. Ruggiero et al. (1986) measured the ESR spectra of samples of lysozyme, myoglobin, and hemoglobin with covalently bound spin labels and noncovalently bound spin probes, in solution and in the partially hydrated powder, over the temperature range 120-260 K. T h e several proteins behaved similarly. The solution samples differed from the powders in showing a change in spectrum shape at 210 K, understood to represent freezing of water in the hydration shell. ESR spectra (Rupley et al., 1980) of lysozyme samples containing a noncovalently bound spin probe, TEMPONE, are strongly dependent

78

JOHN A. RUPLEY AND GIORGIO CARER1

i -k FIG. 20. ESR spectra of TEMPONE noncovalently bound to lysozyme, for hydration levels of 0.02- 1.33 h. The mole ratio of TEMPONE was 0.018; at this low value spin-spin interactions do not make a significant contribution to the measurements. All measurements were made at 24°C. From Rupley et al. (1980).

on hydration level (Fig. 20). The spectrum shifts, over a narrow hydration range, from being characteristic of a motionally restricted solid sample to being characteristic of a solution. The change in motional properties begins sharply at 0.25 h (Fig. 21). This is the hydration level seen in the sorption and heat capacity isotherms as the start of the condensation of water over the weakly interacting regions of the surface. The shape of the spectrum shows a discontinuity at 0.07 h, although the correlation time does not. Thus, both of the discontinuities in the thermodynamics of the interface, seen in the sorption and heat capacity isotherms, appear in the motional behavior of the TEMPONE. Apparently, the same characteristicsof the surface that determine the thermodynamics determine motional properties below 0.38 h. Between 0.25 and 0.38 h the correlation time for the TEMPONE changes from 6 x to 4 x sec. Addition of more water, beyond

79

PROTEIN HYDRATION AND FUNCTION

- *

- 0.4

I

k 0

-

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FIG. 2 1 . Values of the correlation time, T, for TEMPONE noncovalently bound to lysozyme in the variable environment as a function of hydration level. Error bar shows the range of values that gives acceptable simulated spectra. Fraction of TEMPONE in the variable environment is 0.5 2 0.2 at high hydration. From Rupley et al. (1980).

what thermodynamic measurements show is sufficient to obtain the full hydration end point, decreases the correlation time to the solution value of 1O-Io sec.

3 . Comment The results cited above suggest that ESR, NMR, and perhaps other methods that measure motional behavior can detect perturbation of multilayer water, which thermodynamic methods (e.g., sorption or heat capacity) d o not. There are several points that argue against this possibility. (1) T h e motional properties and thermodynamics of a system reflect the same underlying physics. Viewed most simply, a 10-fold change in a rate constant is expected to be associated with a change of 1.4 kcal/mol in barrier height, i.e., in the interactions of a species traversing the barrier. Such a large effect of the protein surface on multilayer water should be detectable by most thermodynamic methods. Motional and thermodynamic properties show parallel changes for hydration levels below monolayer coverage, and one expects that the same should be true

80

JOHN A. RUPLEY AND GIORGIO CARER1

if there were changes above monolayer coverage. (2) A change in motional properties above monolayer coverage may reflect a special chemistry and not a perturbation of multilayer solvent. With regard to the changes in a spin probe correlation time found for lysozyme for hydration levels above 0.38 h, a solute appreciably larger than water should display some motional restriction when constrained to move within the plane of a surface monolayer. Several layers of water may be needed for full solvation of the probe, even though a monolayer may be the only water perturbed by the protein. (3) The models used to interpret resonance experiments may be incomplete. This point is discussed above for NMR. (4)Measurements of hydration forces between surfaces detect perturbations of water that correspond to 100 cal/rnol of water or less for the second layer about a protein. This size perturbation of a barrier height, corresponding to a 25% change in a rate constant, would not be detected by most dynamic measurements of hydration. Collective motions of groups of water molecules perturbed to this extent might be detected, and it is an interesting possibility that resonance methods are monitoring motions of this type.

D . Hydrogen Exchange The rate of exchange with solvent of an amide hydrogen of the polypeptide backbone of a protein has been reduced, as a consequence of the folding, by more than eight orders of magnitude from the rate found for a model peptide, in which the amide proton is fully exposed to solvent (Gregory and Rosenberg, 1986; Rosenberg, 1986). Solid-state measurements of protein amide hydrogen exchange were first made incidentally, in studies of polypeptide structure carried out by infrared analysis of protein films (Haggis, 1956, 1957).The shift in the amide I1 band associated with replacement of the amide hydrogen by deuterium remains a useful method for following the exchange process. Lysozyme has been used often as a model protein in powder (see below) and solution hydrogen-exchange studies (Delepierre et al., 1984; Hvidt and Nielsen, 1966; Woodward and Hilton, 1979). Chirgadze (1972) found that the extent of exchange of a myoglobin film increased with increase in water partial pressure. Hnojewyj (1971, 1978)followed exchange in insulin and hemoglobin powders gravimetrically. Deutschmann and Ullrich (1979), using infrared to monitor the amide I1 band, measured deuterium-hydrogen exchange for lysozyme and other proteins in films. They found that at high humidity the exchange in films was like exchange in solution. Deutschmann and Ullrich (1979) summarized early work on exchange in protein films.

PROTEIN HYDRATION AND FUNCTION

81

Baker et al. (1983) used hydrogen exchange to look for a difference in structure between lysozyme in solution and the dry state. Lyophilized protein was dissolved in tritiated water and exchange in was allowed to proceed for 3 min, after which the solvent was removed on a column and exchange out was followed for several hours. Data obtained in this way for the dry lyophilized protein were compared with data obtained similarly for a solution sample, which was lyophilized protein dissolved 30 min before exchange in. The dry sample showed slower exchange out than the solution sample, with differences of 0.5- 1.5 mol of hydrogen exchanged per mol of protein. This result was understood to reflect a conformation change due to drying, which reversed upon dissolution between 3 and 30 min. Interpretation of the data is clouded by two factors associated with complexities in powder exchange (Schinkel et al., 1985). (1) There is a high salt concentration in lyophilized powders of pH different from the isoionic point; this is a contribution of the counterions, and it is present even if no buffer or salt was added to the solution from which the lyophilized sample was obtained. (2) A nonvolatile acid, such as the sulfuric used to control p H in the experiments by Baker et al. (1983), is concentrated in the powder, to give an effective pH for the powder that is substantially below the nominal pH of the parent solution. Because a protein powder does not dissolve instantaneously, one would see a contribution from the above factors during a short exchange in period of 3 min, as has been observed in other powder dissolution experiments (Schinkel et al., 1985). Both factors would favor rapid exchange of deeply buried protons during the short exchange in, and thus would lead to slower exchange out. The exchange out of tritium from lysozyme powders has been measured for a wide range of water activity (Rupley et d.,1983; Schinkel et al., 1985). Figure 22 shows the number of hydrogens per molecule of lysozyme that remain unexchanged after 24 hr at pH 5, 25°C. The solution exchange rate is reached at about 0.15 h. Apparently, the internal motions monitored by amide hydrogen exchange are independent of hydration above the level where one-third to one-half the surface is covered. This observation is remarkable, considering that motions at the protein surface, as observed in the ESR measurements of TEMPONE dynamics, are unfrozen only above 0.2-0.25 h. One can conclude that motions at the protein surface and the internal motions monitored by hydrogen exchange are not coupled. Hydrogen-exchange rates as a function of level of hydration can be calculated from data such as those of Fig. 22. Results of this kind, for p H 2-10, are given in Fig. 23. The slope of the exchange rate-water

82

JOHN A. RUPLEY AND GIORGIO CARER1

FIG. 22. Hydration dependence of amide hydrogen exchange in lysozyme powder at pH 5. Individual samples of pH 5 fully labeled (with SH20) lysozyme were equilibrated at 25°C for 24 hr at various water contents obtained by isopiestic equilibration (0)or by the addition and mixing of solvent (A).The samples were then dissolved to a concentration of 20 mg/ml and 100-jdaliquots were analyzed by gel filtration. The arrow indicates the 24-hr solution H,, end point. H,, represents the number of hydrogens remaining unexchanged. From Schinkel et al. (1985).

activity profiles is the order in water for the exchange process, which, averaged over all pH, is 2.9 4 0.3. The order in water is independent of pH. At higher pH levels the exchange process senses the more deeply “buried” amides. It is striking that amide protons that show, owing to the folding of the protein, several orders of magnitude difference in exchange rate, show the same dependence of exchange on hydration. This observation suggests that the hydration dependence of arnide exchange for the protein is simply the intrinsic hydration dependence of the exchange process, and the hydration dependence of amide exchange for the protein is the same as would be found for model peptides. If this is true, it is not surprising that the discontinuities seen at 0.07 and 0.25 h in the sorption, heat capacity, ESR, and other measurements, do not appear in the hydrogen-exchange data.

83

PROTEIN HYDRATION AND FUNCTION

-1.0

-0.0

-0.6

-0.4

-0.2

0.C

LOG PI?,

FIG. 23. Dependence on log water activity of log ratio of powder to solution amide hydrogen exchange rate for lysozyme. Log rate ratio data for pH 2 (bottom) to pH 10 (top) are given as a function of log(P/Po).The slopes of the lines give the order of the protein exchange reaction with respect to water. The slopes from least-squares linear regression are the following: pH 2, 2.57; pH 3, 2.90; pH 5, 3.14; pH 7, 3.14; and pH 10, 2.53. Displacement along the log rate ratio axis is arbitrary. Numbers indicate some of the H,, values for which rate ratios were determined. From Schinkel et al. (1985).

Poole and Finney (1983a) measured 8-day out-exchange for deuterated samples of lysozyme as a function of hydration level. Their results are generally similar to those of Schinkel et al. (1985), but detailed comparison is not possible because the samples were not completely deuterated and the pH was not specified. Poole and Finney (1983a) suggested that exposed hydrogens exchange at low humidity, and that a waterinduced increase in flexibility occurs at 0.04-0.07 h, allowing exchange of buried hydrogens. This interpretation is at variance with the data of Schinkel et al. (1985), which show closely similar effects of hydration on exposed and buried hydrogens. Several observations made by Schinkel et al. (1985) in the course of

84

JOHN A. RUPLEY AND GIORGIO CARER1

performing the powder-exchange studies appear to be generally important for measuring and understanding the properties of partially hydrated proteins. (1) Rapid isopiestic equilibration of the powder is possible (half-time, 30 min for a final hydration level 0.15 h) if the sample is thinly distributed and the vapor flux is high, as can be obtained with a large surface area for the solvent reservoir controlling the water partial pressure and essentially no restriction on vapor path between sample and reservoir. (2) T h e half-time for equilibration increases strongly with the final hydration level, as expected from simple kinetic considerations. (3)Jump hydrogen-exchange measurements were used to show that the order of exchange in protein powders is the same as in solution. These results are evidence for similarity of the protein conformation in powder and solution states, but they conflict with the interpretation of Baker et al. (1983). (4) There can be a high effective ionic strength in a partially hydrated powder of low hydration level. Even for a “salt-free” protein, if the system is not isoionic, there will be counterions. For lysozyme at pH 5 and 0.2 h, the effective ionic strength, calculated from the counterion concentration only, is 4.2. A 20 mg/ml solution of this sample would have an ionic strength of 0.016. (5) At low hydration the pH of a powder sample depends strongly on hydration level and is higher than the pH of the solution from which the powder was obtained. This is a result of the high effective ionic strength at low hydration and also of the upward shift in carboxyl pK at low hydration (see Section 11). Points (4) and (5) explain why hydrogen exchange of lysozyme powders under isopiestic equilibration appears to be slightly faster at high hydration levels than exchange in solution (Fig. 22). When water is mixed into the powder at the start of exchange, to give immediately the same final hydration level as that reached slowly by isopiestic equilibration, the exchange rate is the same for high-hydration powders (0.4 h and above) and solutions (Fig. 22). Apparently, during the isopiestic equilibration the transiently high pH and ionic strength in the powder produce rates of exchange sufficiently high for reaction of deeply buried (slowly exchanging) amide hydrogens. E . Spectroscopy

1. Fluorescence

Strambini and Gabellieri ( 1984) found the tryptophan phosphorescence of lysozyme and several other proteins to have similar long lifetimes (about 1 sec) in the dry state. In solution protein phosphorescence lifetimes are generally widely different and short. The long dry-state

PROTEIN HYDRATION AND FUNCTION

85

lifetimes are consistent with reduced mobility of the chromophore environment. At 0.3-0.4 h phosphorescence of partially hydrated protein was similar to that found for solution, suggesting that hydration to this level increases the mobility of the chromophore so that its motion becomes fast on the phosphorescence time scale. Permyakov and Burstein ( 1 977) measured the steady-state fluorescence of tryptophan in several proteins as a function of hydration. They suggested that hydration increases the flexibility of the protein. Sheats and Forster (1983) measured the fluorescence lifetimes of powder samples of bovine serum albumin as a function of hydration. The lifetime increased slightly, from 3.0 to 3.6 nsec, between 0.02 and 0.1 h, changed little between 0.1 and 0.4 h, and increased again at higher hydration, toward the solution value of 6.5 nsec. The hydration dependence of the fluorescence at low hydration is similar to the hydration dependence of amide hydrogen exchange, and it is possible that serum albumin fluorescence similarly reflects principally the effect of water concentration on the underlying chemical event-here, exciplex formation-rather than on the motional or time-average properties of the surrounding protein. In this regard water must diffuse to the chromophore during the lifetime of the excited state, to form the exciplex. Fucaloro and Forster (1985) found substantially different behavior for the hydration dependence of the tryptophan lifetime of chymotrypsinogen A (Fig. 24). Below 0.15 h the lifetime was constant; above 0.15 h the lifetime decreased from the dry protein value of about 3 nsec to the dilute solution value of near 2 nsec at a hydration level above 0.4. T h e change in lifetime near 0.15 h is sharp, as for a phase change. The percolative phase transition of lysozyme is at this hydration level. Azurin has a single buried tryptophan. Fluorescence anisotropy has been measured as a function of hydration level for azurin incorporated in a polymer film (Careri and Gratton, 1986). In the wet film the value of the anisotropy is close to that for azurin in solution at high temperature and low viscosity. At low hydrations and in the dry film, motion of the tryptophan chromophore is frozen. 2. Neutron Scattering Neutron spectroscopy is becoming a principal tool for the study of protein dynamics (Cusack, 1986, 1989; Middendorf, 1984; Middendorf et al., 1984). Current instruments cover motions with characteristic times from lo-’ to 10-13 sec. This range embraces essentially all protein modes excited at room temperature (the soft modes), including motions of the solvent shell and also low-frequency large-scale domain motions, like the hinge-bending motion of the lysozyme domains that form the

86

JOHN A. RUPLEY AND CIORGIO CARER1

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substate cleft. Neutron spectroscopy allows a test of the methodology of protein dynamics simulations, through comparison of measured spectra with those calculated from the results of normal mode analysis (Cusack, 1989; Cusack et al., 1986). Incoherent quasielastic neutron scattering measured as a function of hydration for powders of deuterated phycocyanin has been used to probe water motions (Middendorf et al., 1984). The simplest model accounting for the data was jump diffusion of water molecules between localized-sorption sites and the development of clusters of surface water at higher hydration (half-coverage of the surface, 0.15 h). This model is consistent with the picture developed from sorption thermodynamics.

PROTEIN HYDRATION AND FUNCTION

87

The characteristic jump length was 5-9 A and the residence time 530 nsec. Comparison of neutron scattering of lysozyme at 0.07 and 0.20 h (Smith et al., 1987) showed that hydration decreased elastic scattering and increased inelastic scattering between 0.8 and 4.0 cm-I. This observation is consistent with an increase in the number of low-frequency modes. Normal mode analysis indicates that the lowest frequency mode of lysozyme and the hinge-bending mode fall in this frequency range (Brooks and Karplus, 1985; Bruccoleri et al., 1986; Levitt et al., 1985). Hydration of a protein has little effect on the scattering spectrum, outside of that noted above (Cusack, 1989). Neutron scattering results have been compared with theoretical treatments, specifically normal mode analyses (Cusack, 1989; Cusack et al., 1988; Smith et al., 1989). The experimental data serve as a check on, and have led to improvements in, the assumptions, including the potential functions, of the theoretical treatment. The experimental spectra are relatively smooth, lacking features found in the calculated spectra. Anharmonic motions, sample heterogeneity, and frictional damping, all expected for an experimental sample, favor smoothing of the scattering spectrum. Transitions between conformational substates, which reflect roughness of the potential surface within the envelope of the minimum corresponding to a state of a protein, also are expected to affect the smoothness of the spectrum. Different proteins show similar scattering spectra. This suggests that secondary structural elements, the proportions of which differ widely among proteins, do not dominate the lowfrequency dynamics. Doster et al. (1989) described the temperature dependence of the neutron scattering for myoglobin. They observed the transition near 200 K seen in other dynamical properties. Below this transiton myoglobin behaves as a harmonic solid, dominated by vibrational motions. Near 200 K new degrees of freedom are excited, and a transition is seen in the character of the scattering. Doster et al. (1989) proposed an asymmetric two-state model and a jump mechanism, with torsional degrees of freedom contributing above 200 K. The characteristic time is 0.3-0.5 psec and the characteristic length is 1.5 i% for the fully hydrated protein. T h e size of the effect is smaller for the dry protein. Above 240 K a slower process (i.e., 20 psec) was detected. The protein dynamics and the temperature dependence monitored by neutron scattering are remarkably similar to the dynamics monitored for a longer (i.e., 100 nsec) time scale by Mossbauer spectroscopy. The mean square displacements determined from neutron scattering are about twice those from Mossbauer data.

88

JOHN A. RUPLEY AND GIORGIO CARER1

3. Mossbauer Spectroscopy Mossbauer spectroscopy monitors displacements of 57Featoms that occur in a time shorter than sec. It gives the mean square value of the displacements ((9)) and, for certain modes, the characteristic frequency. The technique is a powerful probe of protein motions (Goldanskii and Krupyanskii, 1989; Parak, 1986, 1987, 1989; Parak and Reinisch, 1986). Proteins such as hemoglobin and myoglobin, with an intrinsic iron atom that can be partially substituted with 57Fe,are clearly suitable for Mossbauer spectroscopic analysis. A Mossbauer label can be introduced into a protein that does not contain iron, such as chymotrypsin or human serum albumin (Belonogova et al., 1979). The Rayleigh scattering of Mossbauer radiation (RSMR), detected by use of an 57Fe-containinganalyzer, gives information on the mean square displacement averaged over all atoms of the sample, and can be used with samples that do not contain 57Fe(Goldanskii and Krupyanskii, 1989; Parak and Reinisch, 1986). Mossbauer spectroscopic measurements have been carried out with the principal aim of understanding protein motion, but they provide valuable insights into the physics of hydration. Mossbauer spectroscopy shows the effect of hydration on the internal motions of proteins. Figure 25 (Parak et al., 1988) compares ( x ’ ) values for dried myoglobin (open squares) and fully hydrated myoglobin (open circles). Below 200 K internal motion is frozen. Above 200 K the dried sample shows about one-half the (x’) value for the heme iron of the hydrated sample. The dependence of (x’) on hydration at 25°C is similar for hemoglobin, myoglobin, and ferredoxin, with a break at 0.5 relative humidity, about 0.15 h; above this hydration level the iron motions increase strongly (Belonogova et al., 1978). Ferritin exhibits a first-order phase transition at 250-280 K, probably peculiar to this protein and associated with a glass transition of the supercooled water about the iron atoms (Bauminger et al., 1987). Mossbauer labels covalently attached to proteins, presumably at the protein surface, exhibit temperature dependences similar to that described above for the heme iron and hydration dependences showing motion developing above a slightly higher hydration level than that found for the heme iron (Belonogova et al., 1979; Likhtenshtein, 1976). Mossbauer spectroscopic measurements suggest that the hydration water of myoglobin and the internal motions of the protein are coupled. [57Fe]Ferricyanidediffused into the solvent of myoglobin crystals exhibits (x2) values equal to those for the heme iron for temperatures below 250 K, and greater than those for the heme iron at higher temperatures (50% greater at 300 K) (Parak, 1986). The [57Fe]ferricyanidein the crystal monitors motions of the hydration water: [57Fe]ferricyanidein “bulk” water shows no Mossbauer spectrum.

89

PROTEIN HYDRATION AND FUNCTION

0.1 0

0.05

100

300

200

FIG. 25. Mean square displacements, {x*), in myoglobin as a function of temperature. X-Ray structure analysis: 0 ,iron; V, histidine (HisFS) bound to the iron; 0 , distal histidine (HisE7); _ _ , linear regression; ---,extrapolation. Mossbauer spectroscopy: 0, deoxymyoglobin; ---, Debye law; , theory; 0 , new experiments with high accufreeze-dried myoglobin. From Parak et al. (1988). racy; 0,

Other measurements also suggest that the hydration water of myoglobin and the internal motions of the protein are coupled. For example, the 10 GHz dielectric response of the water of myoglobin crystals has a temperature dependence close to that of the heme iron (Singh et al., 1981). The 0 - D stretching band (Doster et al., 1986) is also correlated with the above properties (Fig. 26). The temperature dependence of the infrared properties and of the heat capacity (Doster et al., 1986) were interpreted as indicating that the hydration water melts at 190 K and

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90

JOHN A. RUPLEY AND GIORGIO CARER1

that there is a broad glass transition between 180 and 270 K. Suggestions about the mechanism of coupling between solvent and protein can be found in the references cited above. For other evidence bearing on the coupling of solvent and protein motions, see Section VI. Measurements of the Rayleigh scattering of Mossbauer radiation (RSMR) monitor the average motional properties of the protein (Goldanskii and Krupyanskii, 1989; Parak and Reinisch, 1986). Myoglobin RSMR values of (x2> show that at reduced hydration (0.37 relative humidity, -0.1 h) there is no motion at temperatures to 300 K (Krupyanskii et al., 1982) (Fig. 27). Displacements at 0.94 relative humidity and in solution are similar over the full temperature range. The temperature for onset of motion averaged over all atoms of myoglobin is 220-240 K, higher than the temperature for unfreezing of heme iron motion. RSMR measurements made as a function of hydration level for bovine pancreatic trypsin inhibitor, lysozyme, and human serum albumin (Kurinov et al., 1987) show that the response deviates from simple additivity of independent water and protein motions above about 0.1 h. T h e temperature dependence at fixed hydration is like that described for myoglobin. The data are understood to reflect an increase in protein motion as hydration increases from 0.1 to 0.75 h, for bovine pancreatic trypsin inhibitor and human serum albumin. Lysozyme motions are constant above about 0.4 h, and the data of Fig. 27 suggest that this may be true for myoglobin also.

0.L

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FIG. 27. Mean square displacement (x2) averaged over all atoms of myoglobin, but corrected for the water content, determined from Rayleigh scattering of Mossbauer radiation. (Sample a) 0, Lyophilized sperm whale myoglobin hydrated for 3 days at 0.37 relative humidity. (Sample b) 0, Hydrated at 0.94 relative humidity. (Sample c) A , 29.6 wt% solution. (Sample d) 0 ,Myoglobin crystals. From Krupyanskii et al. (1982).

PROTEIN HYDRATION AND FUNCTION

91

A principal point is that the Mossbauer and RSMR signals depend on both the temperature and hydration level of the sample (Goldanskii and Krupyanskii, 1989). For example, in the absence of solvent, there is no motion except that of the harmonic solid at temperatures above 200 K. For additional discussion see Section VI. Pictures of the events associated with the 200 K transition have been given by Parak (1989) and by Goldanskii and Krupyanskii (1989) and are summarized in Section V1. F. Enzyme Activity

The activities of several enzymes have been studied in partially hydrated powders as a function of water activity or water content. Experiments of this type are not difficult to perform. Solutions of substrate and enzyme are mixed quickly, and the mixture is immediately frozen and lyophilized, which stops the reaction and gives a stable dry powder. If appropriately high concentrations of enzyme and substrate are mixed, the powder is of the enzyme-substrate complex. The sample is rehydrated under a controlled atmosphere to give the desired final hydration level. Conditions, particularly the pH of the sample, are set such that the hydration equilibrium is substantially complete (within several hours) before appreciable enzyme reaction has taken place. The problem of defining pH in partially hydrated powders was discussed in Section II,D in connection with hydrogen-exchange measurements. The pH of a powder appears to equal the nominal pH (that of the solution from which the powder was lyophilized) above about 0.15 h.

1 . Chymotrypsin Khurgin et al. (1977) measured the chymotrypsin-catalyzed breakdown of the amide substrate N-succinyl-L-phenylalanine-p-nitroaniline at low hydration levels. For this substrate the acylation process is rate limiting. Figure 28 shows the extent of reaction for 1: 1 enzymesubstrate mixtures, of nominal pH 7.5, reacted for 5-7 days. The intent of the experiments was to define the critical water concentration at which activity could first be detected. This was determined as the intercept of the linear region of the response with the abscissa. For chymotrypsin with no added buffer, the critical hydration level was at relative humidity 0.48, which corresponds to 0.12 h (Luescher-Mattli and Ruegg, 1982a). The reaction grows explosively (Fig. 28) above this hydration level. Addition of 0.57 g of sodium acetate per g of chymotrypsin reduced the critical hydration level by about half. This may reflect the hydration of the the salt, rather than a specific effect on the enzyme. Roslyakov and Khurgin (1971) carried out experiments similar to the

92

JOHN A. RUPLEY AND GIORGIO CARER1

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0.2

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0.4

0.8

PIP, FIG. 28. Hydration dependence of chymotrypsin acylation. Dependence on relative humidity (p/ps) of the extent of conversion of the amide substrate, N-succinyl-Lphenylalanine-p-nitroaniline (SPN), to the nitroaniline product and acyl enzyme, for a 1 : 1 SPN-a-chymotrypsin powder of nominal pH 7.5. Reaction time was 5-7 days. The weight percentages of sodium acetate present in the powder were: curve 1, 0%; curve 2, 6.4%; curve 3, 12%; curve 4, 17%; and curve 5,56.5%. The ordinate, A = D416/Dsj7,is a measure of the nitroaniline product. (From Khurgin et al., 1977.)

above on the deacylation of cinnamoylchymotrypsin, except that all reaction mixtures contained sodium acetate, at concentrations of 0.0630.12 g of salt per gram of chymotrypsin. At the lowest sodium acetate concentration the critical hydration was 0.156 h, and the level decreased with increased salt. The critical hydration level in the absence of salt probably is in excess of 0.2 h. 2 . Lysozyme

Rupley et al. (1980) measured the reaction of lysozyme with the hexasaccharide of N-acetylglucosamine [(GlcNAc),], for pH 8.0- 10.0 at 25"C, over the full hydration range (Fig. 29). High pH was used to slow the enzyme reaction, so that the hydration process was not rate limiting. The effect of pH on the powder reaction followed expectation from solution studies. The threshold hydration level was 0.2 h. Figure 30 shows that the development of enzyme activity closely parallels the development of surface motion, detected by using a nitroxide spin probe (see above). Both processes show 15th-order dependence on the water

93

PROTEIN HYDRATION AND FUNCTION

WEIGHT PERCENT WATER

FIG.29. Enzymatic activity of lysozyme as a function of water content (grams of water per gram of sample), at pH 8, 9, and 10. 0, 0, A , Measurements on powders hydrated by isopiestic equilibration; H, 0 , A, solvent added to powder. Powder samples were the 1 : 1 (GlcNAc)e-lysozyme complex, obtained by lyophilization. The reaction rate ( v o ;sec-l) was determined by product analysis. From Rupley et al. (1980).

0

0

0.1

02

0.3

0.4

0.5

0.6

0.7

0.0

9

g H20/g PROTEIN

FIG. 30. Comparison of ESR and enzyme activity changes with hydration. Effect of hydration on lysozyme dynamic properties. (Curve f ) Log rate of peptide hydrogen exchange. (Curve g) 0, Enzyme activity (log vo); 0, rotational relaxation time (log 7-1) of the ESR probe TEMPONE. From Rupley et al. (1983).

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JOHN A. RUPLEY AND GIORGIO CARER1

activity. The critical hydration level for the enzyme activity is the same as that for protonic percolation on the enzyme-substrate complex (see Section III,A,2). In solution the hexasaccharide is cleaved by lysozyme relatively cleanly to tetramer and dimer. This is true also in the hydrated powder, at hydrations below 40 wt% water. Between 40 wt% water and the dilute solution the pattern undergoes changes, reflecting the contribution of transfer reactions. The reaction rate at full hydration in the powder (i.e., 0.38 h) is about 10% of the solution rate.

3. Other Enzynes Skujins and McLaren (1967)co-lyophilized urease and [ 14C]urea.The rate of reaction, determined by the level of I4CO2,was measured as a function of water content. Onset of enzyme reaction occurred at 0.6 relative humidity. The samples contained a 25 :1 weight ratio of urea to urease. Sorption isotherms measured separately for enzyme and urea showed that below 0.75 relative humidity the urea adsorbed no water, and thus that the enzyme changes reflected adsorption of water by the urease. From the sorption isotherm for urease, 0.6 relative humidity corresponds to 0.15 h. One enzyme reaction has been detected at extremely low hydration. Yagi et al. (1969) found that hydrogenase lyophilized at 10+ mm pressure catalyzed the para-hydrogen-ortho-hydrogen conversion. Stevens and Stevens (1979) measured the hydration dependence of glucose-6-phosphate dehydrogenase, hexokinase, fumarate hydratase (fumarase), and glucose-6-phosphate isomerase (phosphoglucose isomerase) over the range 0.1-0.6 h. Serum albumin was used as a carrier protein to buffer the water content. The hydration isotherms of the enzymes and the serum albumin were assumed to be similar. For the first three enzymes activity was detected (0.05% of full solution activity) near 0.2 h. Activity was measurable for the isomerase at 0.15 h. In all cases, even at 0.3 h, the activity in the powder was less than 5% of the solution rate. Diffusion of substrates in the powder may be rate limiting. The amount of albumin in the powder affected the rate. The food technology literature contains a substantial number of references to enzyme activity at low water content (Acker, 1962; Drapron, 1985; Potthast et al., 19’75).Drapron (1985) gives tables of the hydration level for the onset of activity of various enzymes. Much of this work consisted of monitoring the activity of a particular enzyme, as a function of relative humidity, for a sample (eg., a food product) in which the enzyme was not the principal component. Such measurements have the difficulties of interpretation associated with multicomponent systems-

PROTEIN HYDRATION AND FUNCTION

95

most importantly, the replacement of water by other compounds that might solvate and affect, like water, the properties of the protein. Metabolic activity has been measured in the desiccated state of seeds, spores, and anhydrobiotic organisms such as Artemia (Crowe and Clegg, 1973, 1978; Leopold, 1986). The threshold for metabolic activity is generally 0.2-0.3 h. 4 . Comment

The systematic measurements for chymotrypsin and lysozyme show that these enzymes differ in the hydration level of the onset of enzyme activity. The difference is sufficiently great that experimental artifacts related to the methods of determining the extent and onset of the reaction cannot be the explanation. The other measurements cited above, although perhaps less cleanly interpretable, show threshold hydration levels ranging from below 0.1 to above 0.3 h. Apparently, there is no single hydration level characteristic of the onset of enzyme activity. This is not surprising, because the way in which water of the hydration shell enters into the enzyme reaction should depend on the mechanism of the reaction.

G . Other Measurements

1 . Reverse Micelles, Microemulsions, and Nonaqueous Solvenb Micelles form when a suitable amphiphile [e.g., sodium bis(2-ethylhexy1)sulfosuccinate (AOT)], is introduced into a hydrocarbon solvent (e.g., isooctane). Reverse micelles containing water form when water is taken up by an isooctane-AOT solution. At water contents exceeding what is needed to saturate the polar head groups forming the micelle wall, the system can properly be termed a water-in-oil microemulsion, in which water droplets stabilized by a monolayer of surfactant are dispersed in an organic solvent. For convenience, the terms reverse micelle and microemulsion are sometimes considered equivalent. There is a considerable literature on the properties of proteins, particularly enzyme activity, in reverse micelles (see Luisi and Steinmann-Hofmann, 1987, and references cited therein). The properties of a protein in a reverse micelle depend strongly on water content. Typically, at mole ratios of water to surfactant (w,,) of less than about 3, there is no enzyme activity. As w,, is increased the activity sharply rises, sometimes to an optimum value at w,, = 5-20. The value of for chymotrypsin is as much as 5-fold greater in AOT reverse micelles than it is in aqueous solution (Barbaric and Luisi, 1981; Fletcher

c,,

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JOHN A. RUPLEY AND CIORGIO CARER1

et al., 1985; Martinek et al., 1981). The environment within the micelle is not sufficiently well understood to explain observations of this kind. Measurements of tryptophan phosphorescence of liver alcohol dehydrogenase and alkaline phosphatase in AOT reverse micelles (Gonnelli and Strambini, 1988; Strambini and Gonnelli, 1988) show that the dynamic structures of the macromolewles change over the range of water content, including that of the maximum in the catalytic rate. AOT reverse micelles have been investigated by quasielastic neutron scattering (see Fletcher et al., 1986, 1988, and references cited therein), which showed that at w,, = 20 the water taken up has a diffusion coefficient comparable to that of a high ionic strength aqueous salt solution. Addition of chymotrypsin had no effect on mobility of the surfactant wall. The protein sequestered 5- 10%of the water in the micelle, corresponding to 0.33 ? 0.07 g of water per g of protein. It is not surprising that chymotrypsin activity is detected at low values of w,, ,considering that in powders the onset of chymotrypsin activity is 0.12 g of water per g of protein. The diffusion coefficient of the bound water was reduced 7-fold from the bulk value, comparable to what has been found for a protein in homogeneous aqueous solution. Enzymes are active in organic solvents at low water contents. Porcine pancreatic lipase in glycerin tributyrate (tributyrin) shows, for 0.0 15% water in the tributyrin-pentanol reaction mixture, a rate of transesterification comparable to the value in aqueous solution (Klibanov, 1986; Zaks and Klibanov, 1984). The water content of the protein in the above reaction mixture was 0.01-0.03 h. This is below the level expected for the onset of enzyme activity in protein-water powders. Nonaqueous solvents can produce change in the substrate specificity of an enzyme (Zaks and Klibanov, 1986; Zaks and Klibanov, 1988a) and possibly can lock the enzyme into a more active conformation (Russell and Klibanov, 1988). The dependence of the catalytic activity on added water has been measured for several enzymes in several solvents (Zaks and Klibanov, 1988b). Interesting chemistry is associated with micellar and nonaqueous environments. Three-component systems, however, can be difficult to understand, and for the present our knowledge of protein hydration in twocomponent systems is more likely to throw light on three-component systems than the reverse. 2. Viscosity

Change in solvent viscosity has been found to alter the dynamics of ligand binding (Beece et al., 1980) and enzyme catalysis (Gavish and Werber, 1979). These effects were interpreted in terms of the Kramers

PROTEIN HYDRATION AND FUNCTION

97

theory of reaction rates, which is based on Brownian diffusion of reacting elements over a potential barrier, and explicitly includes a frictional dissipative term that incorporates the viscosity of the medium. In the Kramers regime the reaction rate should vary inversely with the solvent viscosity. Solvent viscosities from near that of water to several orders of magnitude higher were obtained by a change in temperature, the addition of alcoholic or polyhydric cosolutes, or both. The protein rate processes followed a power law in viscosity, with exponent from - 0.5 to - 1, in accord with modified Kramers theory. The rates became independent of viscosity at very high solvent viscosities. At very low solvent viscosities the Kramers relationship should also fail. As shown above, enzyme catalysis and protein rate processes appear to be very slow in the dry state, where the environment is a vacuum and has zero viscosity, for which the Kramers relationship would predict, incorrectly, a rate of reaction higher than that in water. Immobility in the dry state likely has nothing to do with the medium viscosity, however, but rather follows from the absence of the plasticizing action of water. Small amounts of water should function as a plasticizer, catalyzing conformational transitions through affording alternative hydrogen-bonding arrangements (Chirgadze and Ovsepyan, 1972a). The effective viscosity of the solvent at the protein surface is likely greater than the bulk solvent viscosity. The diffusion constant of water at the protein surface is five times smaller than the bulk water value (Polnaszek and Bryant, 1984a).This effect probably can be neglected in experiments such as those discussed in the previous paragraphs, which cover several orders of magnitude in solvent viscosity. The introduction of solvent into molecular dynamics simulations of proteins produces complex changes in motional properties (Brooks and Karplus, 1986) and generally decreases the time constants of atom and group motions (Ahlstroem et al., 1987; Levitt and Sharon, 1988).This is in qualitative agreement with experiments, in that addition of water unfreezes both surface and internal motions of groups of atoms. Amide hydrogen exchange displays curious chemistry. There is an effect of solvent viscosity (Rosenberg, 1986; Rosenberg and Somogyi, 1986; Somogyi et al., 1988), like that for the myoglobin-oxygen reaction. The fastest-exchanging hydrogens show a Kramers-type viscosity dependence. The slowly exchanging amide hydrogens, presumed to be those most buried by the folding of the protein, appear to be affected by the solute, glycerol, used to change the solvent viscosity; that is, the exchange rate reflects the effect of the cosolvent on the unfolding equilibria of the protein. For protein powders, on the other hand, the exchange

98

JOHN A. RUPLEY AND GIORGIO CARER1

rate for both fast- and slow-exchangingamide protons reaches the dilute solution value at below half-coverage of the surface with water. Apparently, in the protein powders the internal motions that are sensed by hydrogen exchange are uncoupled from the surface and its hydration. The exchange events in the partially hydrated powder appear to be identical in rate and character to the events in dilute solution, despite a substantial difference in surface environment. It is not clear how these powder results relate to the general viscosity effect and specific solute effect found for solution exchange in high-glycerol solvents. Somogyi et al. (1988) measured the rate of isotope exchange at the ring nitrogen of Trp-63 of lysozyme as a function of solution viscosity. The data were described by a modified Kramers relationship, with viscosity exponent 0.6. This is similar to what was found for the fastexchanging amide protons of lysozyme. Both processes are of low activation energy and are expected to be subject to viscous damping. 3 . Mechanical Properties Morozova and Morozov ( 1982) measured the viscoelasticity of crystals of triclinic lysozyme and its complex with the substrate analog (GlcNAc)s [the p( 1+4)-linked trisaccharide of N-acetylglucosamine],as a function of water partial pressure. The data are consistent with a model in which two rigid domains are connected by a flexible link. The compliance decreased with decreasing relative humidity, to a limiting low value at 0.4 relative humidity. Binding of the substrate analog reduced compliance. The spring constant estimated for the model was close to the value calculated by McCammon et al. (1976) for the hinge-bending mode of lysozyme. It is possible that this motion is frozen at 0.4 relative humidity. Morozov and Gevorkyan (1985) observed a temperature-dependent change in Young’s modulus, centered at 200 K, which they call a mechanical glass transition. The magnitude of the effect decreases with decreasing hydration. A transition at 200 K has been observed with other dynamic measurements. At 25°C Young’s modulus of lysozyme crystals increases slightly as the hydration is decreased from 0.4 to 0.2 h; it increases sharply below 0.2 h (Morozov et al., 1988), the hydration level at which various other properties of the partially hydrated protein (e.g., heat capacity) show sharp changes. Baer, Hiltner, and colleagues (see Hiltner, 1979, and references cited therein) have used dynamic mechanical analysis to examine the hydration of collagen, elastin, and several polypeptides. A torsional pendulum constructed of the sample was examined for low-frequency (i.e., 1Hz) mechanical loss as a function of hydration and temperature. A common feature is a dispersion that is absent in the dry protein and appears at

PROTEIN HYDRATION AND FUNCTION

99

very low hydration levels (i.e., 0.01 h) and temperatures near 200 K. With increasing hydration the intensity of the response increases and the characteristic temperature decreases to near 150 K. IV. STRUCTURE A . Dafraction

The flow of structural information on the solvent around proteins coming from X-ray and neutron diffraction analyses has increased enormously. There are excellent reviews of this work (see, e.g., Baker and Hubbard, 1984; Edsall and McKenzie, 1983; Finney, 1979; Kossiakoff, 1983; Saenger, 1987; Savage, 1986a; Savage and Wlodawer, 1986; Schoenborn, 1984). Edsall and McKenzie (1983) gave highly useful descriptions and tabulations of crystallographic results for individual proteins. Several recent surveys of the diffraction literature center on aspects of protein hydration: the distributions of water around the 20 different amino acid residues (Thanki et al., 1988), hydration of helices (Karle and Balaram, 1989; Sundaralingam and Sekharudu, 1989; Sundaralingam et al., 1987), and helix geometry (Barlow and Thornton, 1988). Considering the abundance of reviews, this discussion is focused on the results for one protein, lysozyme, for which there is a thorough analysis and categorization of the crystallographic picture of the solvent. Shorter descriptions are given of recent results with several other proteins. These and other analyses indicate that the picture developed for lysozyme is typical of globular proteins.

1 . Lysozyne Blake et al. (1983) refined the structures of human lysozyme (HL) and tortoise egg white lysozyme (TEWL) to 1.5 and 1.6 A resolution, respectively. The diffraction was modeled as arising from three components: the protein, ordered water, and disordered water. Most of the water in the crystals (i.e., SO-SO%) is disordered. The analysis located 143 molecules of ordered water out of about 350 per HL molecule, and 122 molecules out of 650 per TEWL molecule. The ordered water covers 75% of the available surface of the the protein. One-third (TEWL) to one-half (HL) of the total surface is unavailable for analysis of the adjacent water, owing to crystal contacts or disorder in the protein region. Thus, the estimate of surface coverage is in good agreement with the 300 molecules of water estimated by heat capacity measurements as full hydration (0.38 h). The area covered per water molecule is estimated as 18.9 A2

100

JOHN A. RUPLEY AND GIORGIO CARER1

for HL and 2 1.4 hi2 for TEWL. This value is in agreement with powder hydration measurements: the area covered per water molecule, calculated from the heat capacity end point, is 20 A2 (Yang and Rupley, 1979). The ordered water is in a monolayer about the protein. No ordered water is more than about 4.5 A from the surface. The average number of hydrogen-bonded neighbors is two to three. If the interaction between water in the monolayer and the bulk solvent is taken into account, then, on average, there should be nearly one more hydrogen bond, in a direction roughly orthogonal to the protein surface. Hence, the number of hydrogen-bonded neighbors for water in the surface monolayer is similar to the average for bulk water. In view of the multiple hydrogen bonding of the surface water, networks, in the sense of extended chains or clusters of hydrogen-bonded water and protein atoms, should be typical of the hydration shell. Watenpaugh et al. (1978) described extensive water networks for rubredoxin, the first protein for which water arrangements were described. Teeter (1984) found pentagonal closed arrays of water in the crambin crystal. James and Sielecki (1983) described a pentagonal water cluster in penicillopepsin. Regular structures of this kind are found in clathrates and also in simulations of water and aqueous solutions. Icelike and other regular structures, to be distinguished from threads or clusters of hydrogen-bonded waters, were not reported for lysozyme. Analysis of the Debye-Waller B factors suggests that 33-35 waters are strongly bound. These are located mostly at positions that are equivalent in the HL and TEWL structures. Hagler and Moult (1978) noted the similarity in water positions determined for two crystal forms, triclinic and tetragonal, of hen egg white lysozyme. The waters found for the hen egg white proteins are also largely equivalent to ones found for HL and TEWL. These observations suggest that essential features of the water structure in the crystal are intrinsic properties of the hydrated protein and would be found also in the solution state. Tables IV-VI, from Blake et al. (1983), describe the contacts made by the ordered water. The waters are nearly all bonded to polar protein atoms. The infrequent interaction with amide NH reflects the inaccessibility of these atoms. Charged side chains generally bind two ordered waters; polar groups, one. Kundrot and Richards (1987, 1988) described the solvation shell in the hen egg white lysozyme crystal, in connection with a study of the compressibility of protein and solvent. Mason et al. (1984) carried out a neutron diffraction analysis of triclinic lysozyme at 1.4 hi resolution, with 239 water molecules included in the refinement.

101

PROTEIN HYDRATION AND FUNCTION

TABLE IV Ordered WaterMohcuhs about Lysozyme a Number of water molecules Category

HL

TEWL

Asymmetric unit of crystal Making two or more hydrogen bonds to protein Making hydrogen bonds to two protein molecules Making single hydrogen bonds to protein Making hydrogen bonds to other bound water molecules, but not to protein Making no hydrogen bonds to either protein or water molecules Too close (<2.6 A) to protein molecules Total number of water molecules associated with a single protein molecule [(b) + 2(c) + ( d ) + ( e ) ]

143 35 13 44

122 33 12 52

35

19

12 4

2 4

140

128

"HL, Human lysozyme; TEWL, tortoise egg white lysozyme. (From Blake et al., 1983.)

Kachalova et al. (1987) measured the effect of dehydration on the diffraction of three crystal forms of lysozyme, at 3 or 6 hi resolution. The lowest relative humidity studied was 0.4. The molecules as a whole were displaced on dehydration by 1.9-5.2 hi, through rotation and translation of the units. There was also displacement of the two domains of lysozyme relative to one another. The domain displacements of 1.4-

TABLE V Protein- Water Interactions in Two Lysozyme Speciesa

Parameter

.

Number of water molecules hydrogenbonded to protein % Bound to peptide CO % Bound to peptide NH % Bound to side chains % Bound to protein oxygen % Bound to protein nitrogen Mean H20-0 distance (A) Mean HzO-N distance (A) Mean H 2 0 - H 2 0 distance (A)

HL

TEWL

105 42 18 40 68 32 2.83 2 0.15 2.96 f 0.12 2.82 2 0.23

109 44 18 38 64 36 2.81 0.15 2.96 2 0.17 2.84 ? 0.23

*

&HL,Human lysozyme; TEWL, tortoise egg white lysozyme. (From Blake et al., 1983.)

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JOHN A. RUPLEY AND GIORGIO CARER1

TABLE VI Hydration of Side Chains of Lysozyme" HL Side chain

ASP LYS Asn Glu Arg Thr TYr Ser Gln TrP His Cod NH*

Total number 8 6 10 3 14 5 6 6 6 5 1

130 130

Number ordered

TEWL Waterfside chain

Total number

6 5 8 2

2.0 1.8 1.6

-

1.5

9c

6

1.5 1.2 1 .o 0.7 0.7 0.2 0.50 0.24

8 8 6 8

4 4 6 4 5 0 128 129

Number ordered

16b

116

9

6

5 2 130 130

-

5c 5

7 5

7 5 2 124 129

Waterkide chain 1.6b 1.4 1.3c 1.6 1 .o 1.2 0.8

0.2 0.5 0.54 0.24

OHL, Human lysozyme; TEWL, tortoise egg white lysozyme; -, not applicable. (From Blake ct al., 1983.) bCombines Asp and Asn. 'Combines Glu and Gln. dMain-chain peptide groups.

3.6 8, were principally rotations. The crystal forms differed in the extent of the displacements and in the balance between rotation and translation. 2. Crambin

x

Crambin is a 46-residue hydrophobic plant protein. It crystallizes from ethanol-water mixtures. Exceptional1 high-resolution X-ray difresolution data at 300 K fraction data have been obtained [0.945 (Hendrickson and Teeter, 1981) and 0.83 A data at 140 K (Teeter and Hope, 1986)l. Neutron diffraction data were obtained at 1.5 h; resolution and 300 K (Teeter and Kossiakoff, 1984). These results were summarized and discussed by Teeter and Whitlow (1986). The diffraction analyses located 73 of the approximately 90 waters per crambin molecule in the crystal. The water is highly ordered. Nearly all of the water is part of a single hydrogen-bonded network. Much of this water is arranged like that of lysozyme, in chains interacting with polar or charged groups at the protein surface. Some of the network is disordered, in that it consists of chains of waters with alternate positions. A special feature

PROTEIN HYDRATION AND FUNCTION

103

of the crambin crystal is a cluster of five linked pentagonal rings of hydrogen-bonded waters. 3. Myoglobin

Parak and collaborators (Hartmann et al., 1987; Parak et al., 1987) carried out X-ray structure analyses for metmyoglobin at temperatures of 80-300 K. One hundred sixty water molecules, more than one-third of the water in the crystals, were included in the refinement. The disorder is higher for water that for protein atoms (Fig. 31). Reduction in temperature partially freezes out the disorder for the water, as for the protein. The residual disorder at low temperature has been understood to represent conformational substates or a distribution of conformations, frozen in at low temperature and in mobile equilibrium at high temperature. A neutron diffraction analysis of oxymyoglobin (Phillips, 1984) located 120 water molecules at the protein surface. Peters and Peters (1986) analyzed these and earlier X-ray results (Hanson and Schoenborn, 1981; Takano, 1977a,b) and described networks formed of protein polar groups and water molecules. The network structures found for myoglobin appear to be like those for other globular proteins. m

I

I . . . . . . 200

100

300

FIG. 31. Averaged mean square displacements for water and protein atoms of myoglobin. 0 ,All backbone NCCO atoms; 0, all side-chain atoms; *, 160 water molecules found crystallographically;, linear regression; linear regression, neglecting the values at 300 K; ---,extrapolations. From Parak el al. (1987). -0-,

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JOHN A. RUPLEY AND GIORGIO CARER1

4 . Insulin

Baker et al. (1987,1988)described a 1.5 A resolution structure of 2Zninsulin. They located 282 of the estimated 285 waters per insulin dimer in the crystal. These were distributed among 349 sites: 2 17 of occupancy 1.0;126 of occupancy 0.5; five of occupancy 0.33;and one of occnpancy 0.25. There was evidence for ordered water at a distance 8 8, from the protein surface. Nearly 100 waters were bonded only to other waters. The extent of order of the water, judged by B values, increased with an increased number of interactions with the protein. The waters bonded to the protein act as anchors for chains of less well-ordered waters, which are often linked by threads of density, possibly representing paths along which the less-ordered waters are found. There were alternate water positions, sometimes collected into networks of partially occupied sites. Cyclic water structures were found. The protein-water contacts showed preferred geometries. Baker et al. (1988)gave particularly elegant descriptions of the crystal water. 2Zn-insulin is like crambin and rubredoxin in that the crystal has a low proportion of water-for insulin 30.6% by weight. These proteins are similar also in having much of the crystal water ordered sufficiently for detection by X-ray diffraction analysis. Caspar et al. (1988)described an elegant analysis of the diffuse X-ray scattering from insulin crystals. They found two types of coupled motion: one with a characteristic length of about 6 8, and amplitude of about 0.4 A, the other with a characteristic length of about 20 and smaller amplitude. The latter motion represents the jiggle of neighboring molecules of the lattice. The former represents the coupled fluidlike fluctuations within a protein molecule. The short-range motions appear to be similar to those detected by Mossbauer spectroscopy. 5 . Other Proteins Water arrangements essentially similar to those for lysozyme have been found for various other high-resolution structures of proteins, for example, penicillopepsin (James and Sielecki, 1983),ferricytochrome c' (Finzel et al., 1985), glyceraldehyde-3-phosphate dehydrogenase (Skarzynski et al., 1987),and bacteriophage T4 lysozyme (Weaver and Matthews, 1987).Wlodawer et al. (1988)described the solvent about phosphate-free ribonuclease A, at 1.26 A, and have compared their results with those for other high-resolution structures of this protein.

6. Actave-Site Waters

The solvation of the active-site cleft of lysozyme (Blake et al., 1983) does not seem to be different from that of the rest of the protein surface.

PROTEIN HYDRATION AND FUNCTION

105

Nevertheless, there are interesting features of the water arrangements in the cleft. There are water molecules hydrogen bonded to the protein groups that, in the enzyme-substrate complex, form hydrogen bonds to substrate, and the water is at the position that would be occupied by the substrate atom. One water may mediate an enzyme-substrate bond. One or mme waters may be trapped by the hound substrate. A water molecuk bonded to Glu-35 may not be dispIaced by the bound substrate and may be in a position to attack the glycosidic bond. Blevins and Tulinsky (1985) made the suggestion, based on a 1.67 8, resolution structure for a-chymotrypsin, that several molecules of the specificity-site water are not displaced on substrate binding and may serve to position the substrate. Quiocho et al. (1989) analyzed the role of water in the binding of substrate to L-arabinose-binding protein. Despite the considerable difference in structure, D-galactose is bound nearly as strongly as L-arabinose, and D-fucose, only slightly less strongly. Ordered water, which is part of the active site and provides mediating hydrogen bonds between substrate and protein, facilitates the adjustments required to accommodate the various saccharides. The D-galactose-binding protein, which lacks the equivalent of the water molelcules of the L-arabinose-binding protein, does not bind D-fUCOSe or L-arabinose. The kinetics of the binding of inhibitors to thermolysin appear to be controlled by the displacement of a water molecule from the active site (Holden et al., 1987). Meyer et al. (1988) gave a detailed discussion of the possible roles of water in the catalytic mechanism of pancreatic elastase. These include (1) reorganization of solvent about substrate and enzyme during binding; (2) participation of water in bond rearrangement, through enhanced polarization of the catalytic groups and through connecting the catalytic groups to the surface by a chain of hydrogen-bonded waters in a tunnel structure; and (3) homology of this and other water structures among the trypsinlike proteases. A water molecule is bound in the oxyanion hole of proteinase K, a member of the subtilisin family (Betzel et al., 1988). Its replacement by the carbonyl oxygen of the substrate is suggested to be concerted with formation of the tetrahedral intermediate. Oefner and Suck (1986) proposed that a water bonded to histidine is the nucleophile in the hydrolysis catalyzed by DNase I, and that it is part of a Glu-His-water triad reminiscent of the familiar Asp-His-Ser triad of the serine proteases. A water molecule held between two carboxyl groups is possibly the nucleophile in the reaction catalyzed by the aspartic proteinase from Rhizofm chinemis (Suguna et al., 1987).

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JOHN A. RUPLEY AND GIORGIO CARER1

Wright (1987) found five clusters of more than three water molecules in a 1.8 A resolution analysis of the wheat germ agglutinin dimer. The four saccharide binding sites of the dimer showed similarity in ordered water structure. Karplus and Schulz (1987) found a cluster of 104 water molecules at the dimer interface of glutathione reductase. Brown et al. (1987) compared the structures of wild-type tyrosinetRNA ligase (tyrosyl-tRNAsynthetase) and the Thr-5 1+Pro-5 1 mutant. The stronger affinity of the mutant for tyrosyl-adenylate complexes was attributed to the absence for the mutant of an unfavorable desolvation of residue 51. A halobacterial ferredoxin has a domain rich in aspartate and glutamate that extends from the main body of the molecule (Sussman et al., 1989).The surface charge distribution may increase hydration and may be necessary for function in a saline environment. Reports of high-resolution structures now can be expected to contain a description of a substantial number of ordered waters, some of which are likely to be part of an active site. The above citations are perhaps typical, and are not intended to be exhaustive. 7. High-Resolution Analyses

Savage (1986a-c; Savage and Wlodawer, 1986) summarized descriptions of water structures in ice and hydrates of small molecules, for which there are more results at higher resolution, below 1.O A, than are available for proteins. The preferred geometries found for interactions of water with compounds such as coenzyme B12serve as a basis for modeling protein-water interactions. Savage presented a thoughtful discussion of the problems and procedures for the analysis of solvent in protein crystals. 8. Comment

Structural information and tabulations of the type given for lysozyme are of the greatest importance in understanding hydration. Nevertheless, the consumer of results of analyses of ordered water in protein crystals perhaps should keep in mind several caveats: 1. Most of the ordered water is not “strongly ordered”; that is, site occupancy is less than unity, thermal motion is high, or both. It may be that a different set of water sites can satisfactorily fit the diffraction data. This possibility is discussed by Savage and Wlodawer (1986), who noted that alternative solvent networks were found for crystals of coenzyme B,2- Fluctuations in structure are expected, considering the physics of water.

PROTEIN HYDRATION AND FUNCTION

107

2. The identification of full occupancy and low thermal motion with strong binding may be misleading. Few, if any, water molecules remain in place for as long as the characteristic time for rotational motion of the protein. The free-energy difference between the hydration shell and the bulk solvent is, on average, small (i.e., less than the ambient thermal energy). The interaction of water at the protein surface is not qualitatively different from the interaction of water with other waters. If one could stand on a molecule of bulk water and survey the surroundings, as one figuratively can stand on a protein molecule in the crystal, then the tetrahedrally arranged sites neighboring the water would be seen to be filled with high occupancy, as are the surface sites of the protein. B . Spectroscopy

1 . Infrared (IR) and Raman

The IR and Raman spectra of partially hydrated proteins are a rich source of fundamental information on both water and protein species, owing to the sensitivity of vibrational modes to hydrogen bonding. The similar chemistry of water-water and water-peptide interactions requires that there be great accuracy in spectroscopic measurements of the hydration process. Since the review of the field by Kuntz and Kauzmann (1974), the Fourier transform technique for IR and the tunable laser for Raman spectroscopy have offered important improvements in methodology. Careri et al. (1979b) studied the stepwise hydration of lysozyme films, using differential IR spectroscopy to follow the amide I frequency shift (1660 cm-') (Figs. 32 and 33) and the absorbance at the carboxylate (1580 cm-l) (Fig. 33) and OD stretching (2570 cm-l) frequencies. The apparatus monitored sample weight concurrently with the IR spectrum. The principal conclusions from this work are: (1) the hydration process first affects the carboxylic side chains, producing carboxylate species, detected at 1580- 1600 cm-I; about 40 mol of water per mol of lysozyme interacts with carboxylate groups in the first stage of the hydration process. (2)Weaker water binding sites, mostly amide carbonyl groups of the backbone, are progressively hydrated and accommodate about 100 mol of water per mol of protein. (3) At higher hydration condensation of water occurs to complete the hydration process. (4) No waterinduced conformation changes were detected in the frequency range investigated. The changes in the amide I band with increased hydration are consistent with the expected effect of hydrogen bonding of the amide carbonyl group. (5) Several details of the absorption bands revealed a transition in the water-protein system, close to 0.06 h. The shape of

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JOHN A. RUPLEY AND GIORGIO CARER1

0.10

i

n

_,h

= 0.352

1900

I? I0 Y

(cm-')

FIG.32. IR spectrum of dry lysozyme and difference spectrum for 0.07 h at 27°C. (Top) Dry sample; (bottom) difference spectrum at 0.07 h. From Careri et al. (1979b).

the OD bending band showed the existence of different types of adsorbed water. (6) The sorption isotherms were fit closely by a model based on two classes of primary sorption sites, in agreement with the spectroscopic data (Fig. 33). Careri et al. (1980) and Rupley et al. (1983) compared the above data with other time-average and dynamic properties, to develop a picture of the hydration process. Poole and Finney (1983b),using direct difference IR and laser Raman spectroscopy, studied the hydration of lysozyme. On the basis of these measurements, they suggested that there are conformational changes just below the hydration level for the onset of enzyme activity (i.e., 0.20.25 h). This conclusion conflicts with that of Careri et al. (1979b). Poole and Finney (1984) extended these measurements to lactalbumin.

PROTEIN HYDRATION A N D FUNCTION

109

c

0

8

h

g

100

I

E

.-

x

0

E

b 50

8 0

0.10

0.20

0.30

h

FIG.33. Changes in the carboxylate and amide IR bands with hydration. Shown are the

IR absorption of the carboxylate band at 1580 cm-1 in the difference spectrum (0)and the sum of the absolute value of the absorption of the amide I' difference band at 1645 and 1690 cm-I ( x ) . Data are shown for 38°C (top) and 27°C (bottom). Curves A, D,O sorption onto strong sorption sites; curves B, D,O sorption onto weak sorption sites; curve C, D20 multimolecular adsorption. Curves A-C were derived from the sorption isotherm by fitting data to a three-site model. Ordinate units represent percentages of the values at 0.33 h. From Careri et al. (1979b).

Poole and Barlow (1986), by monitoring the desorption process with IR measurements, studied water removal from ion-paired acidic groups. By the same method Barlow and Poole (1987) measured the strength of water binding to the carbonyl groups in a-helical and p-sheet proteins and found that the strength of water binding to the CO groups is lower in P-sheet than in a-helical proteins. Jakobsen et al. (1986) used Fourier transform IR techniques to study the amide I band for serum albumin in saline solution and in films of a varied hydration level. They concluded that the bandwidth of the amide I is directly related to the amount of adsorbed water and that a more ordered form of helix secondary structure is produced as tightly adsorbed water is removed from the albumin. Doster et al. (1986) measured the OD stretching bands of water in

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JOHN A. RUPLEY AND GIORGIO CARER1

myoglobin films at a varied hydration level, between 100 and 300 K. There was a broad glass transition, centered on a temperature that increased with increasing hydration. Below 190 K the hydration water is frozen. The specific heat of water in myoglobin crystals was determined in parallel experiments; it showed a glass temperature near 220 K. The IR data suggested that the transition could be described as the melting of “amorphous ice” and that this solvent network is composed of water clusters with relatively strong internal bonding. Doster et al. (1986) used this information to address the problem of dynamic coupling of solvent motions with-internal protein motions, suggesting that the cooperativity of the solvent network provides the coupling mechanism. Vander Meulen and Ressler (1980) measured the near-IR spectra of proteins in aqueous solution and compared them with the spectra of protein films. Brown et al. (1983) reported multiple internal reflectance spectra of hydrated films of carbonmonoxy and oxy forms of hemoglobin. This work was extended by Findsen et al. (1986), who, using resonance Raman scattering, measured the effects of hydration on the equilibrium and dynamic properties of hemoglobin and its carbonmonoxy complex. There was a substantial effect of hydration on the CO vibration, but no significant effect on the vibrational properties of the heme protein. Ataka and Tanaka (1979) measured both the far-infrared and Raman peaks of crystalline lysozyme, with self-consistent results. They assigned the uniform background to residual water molecules. Cavatorta et al. (1976)studied, by laser Raman spectroscopy, the intensities of CH and OH stretching for aqueous solutions of lysozyme at different concentrations and explained their findings in terms of a water model with three water species, each with different Raman intensities. Samanta and Walrafen (1978) have repeated and extended this study. They obtained somewhat different data and offered an interpretation involving only a single interaction-namely, hydrogen bonding between lysozyme and water. Aliotta et al. (1981) improved their Raman technique by using isotopic substitution and depolarization, and concluded that lysozyme does not perturb the OH stretching of water. 2 . Absorbance

Lyophilized cytochrome c, when dissolved, shows a change in absorbance, with a half-time of about 1 min (Aviram and Schejter, 1972).The character of the spectrum change suggested that the conformation difference between dry and solution states is the replacement of Lys-79 by Met-80 as ligand to the iron. Giannini and Gratton (1977) measured the ultraviolet spectrum of ly-

PROTEIN HYDRATION AND FUNCTION

111

sozyme as a function of hydration level. Below 0.25 h the tyrosine spectrum was hydration dependent; above 0.25 h it was constant. 3 . Circular Dichroism

Chirgadze and Ovsepyan ( 1972b) measured circular dichroism in the range of 200-240 nm for films of several proteins at varied humidity. Comparison of the spectra with measurements for aqueous solutions showed that the native structure of globular proteins is almost completely retained at high humidity and slightly deformed at low or medium humidity. Figure 34 shows spectra for lysozyme and ribonuclease. The circular dichroism spectra of proteins are determined by more than the secondary structure of the macromolecule. In the case of lysozyme, the tryptophan chromophores contribute (Tanaka et al., 1975).This and other effects may explain the qualitatively different effect of hydration on the spectrum at 220 and 210 nm, seen in Fig. 34. A comparison of conformations based on circular dichroism should be viewed as having substantial uncertainty.

h.nm

FIG. 34. Circular dichroism spectra of (a) lysozyme and (b) ribonuclease in the solid state (films on quartz plate) at various humidities and in neutral aqueous solution. The figures accompanying the curve refer to relative humidity in percentage. In the case of lysozyme, the curve for the solution is identical in shape to the curve of the film at 96% humidity. From Chirgadze and Ovsepyan (1972b).

112

JOHN A. RUPLEY AND GIORGIO CARER1

4 . Magnetic Susceptibility

The diamagnetic susceptibility is a measure of the averaged electronic distribution in bulk matter. Careri et al. (1977, 1980) showed that the differential diamagnetic susceptibility per gram of water adsorbed on lysozyme powders reached the bulk water value at 0.2 h. Lysozyme behaved as a normal diamagnetic substance. The diamagnetic susceptibility and the enthalpy of sorption for lysozyme change similarly at low hydration. Lumry et al. (1962) measured the hydration dependence of the magnetic susceptibility of cytochrome c. Cytochrome c dried by lyophilization differed in susceptibility from protein dried by dehydration.

V. COMPUTER SIMULATION A . Molecular Dynamics A recent review (Brooks and Karplus, 1986) describes methods of including solvent in protein dynamics computations. Simulations of a neon-water system (Geiger et al., 1979) and an alanine dipeptide-water system (Karplus and Rossky, 1980; Rossky and Karplus, 1979) showed that the solvent adjacent to the solute is slightly but significantly perturbed. For the alanine dipeptide the first-layer water had two to three times slower diffusive and rotational motion, and a small (i.e., less than 0.1 kcal/mol) change in the mean hydrogen bond energy. There was only a small change in the number of hydrogenbonded neighbors for the first-layer water, but the total number of neighbors was one fewer than for bulk water. This difference is a reflection of a restricted orientation of the first-layer water, which presumably is at the heart of thermodynamic and other differences between the firstlayer water and the bulk solvent. Post et al. (1986) carried out a molecular dynamics simulation for hen egg white lysozyme, with and without bound substrate. The 53 water molecules most strongly localized in the crystal structure were included in the computation. The dynamics of the water were similar to what had been inferred from the diffraction data through analysis of the DebyeWaller B values for the solvent and adjacent protein atoms. In a simulation of the active-site region, Brooks and Karplus (1986, 1989) found that water molecules formed networks that stabilized charged residues, particularly neighboring groups of like charge. They suggested that there is a possible functional role for collections of like-charge groups and the water networks associated with them. The magnitude of the

PROTEIN HYDRATION AND FUNCTION

113

solvent effect on dynamic properties of the protein depends on there being spatial coupling of protein to solvent atoms, and on there being dynamic coupling (i.e., overlap between the time scales of protein and solvent motions). T h e structure and dynamics of the solvent are similar to what has been observed for the solvent about small molecules. T h e range of environments of the solvent atoms is reflected in the geometry and dynamics of the solvent. For example, water mobility is reduced about charged groups. It may be significant that solvent characteristics are influenced by distant protein groups. Ahlstroem et ul. (1987) described simulations for parvalbumin in uucuo and in a system with 2327 water molecules and three sodium ions for electroneutrality. The simulation with water differed less from the crystal structure than the in uucuo simulation. Large effects on dynamics were found for protein interior as well as surface atoms (Fig. 35). Levitt and Sharon (1988) carried out a 210-psec simulation of fully solvated bovine pancreatic trypsin inhibitor (BPTI) with 2607 water molecules. The periodic boundary was a rectangular box 8 hi larger than the protein. All hydrogen atoms and all degrees of freedom were included in the computation. Carboxyl and amino, but not guanido, groups were charged. The solution simulation equilibrated two times faster and showed about one-half the root mean square (rms) deviation from the X-ray structure, compared with an in uucuo simulation. Fluctuations are about 30% smaller in the solution simulation. All mainchain hydrogen bonds found in the solution simulation are found in the X-ray structure; one X-ray structure bond is missing in the simulation. The hydrogen bonding of the in uucuo simulation shows substantial deviation from the X-ray structure. Clearly, the solvent has an effect on the properties of the protein. Addition of solvent eliminates the protein-vacuum interface and thus the shrinking of the protein found in uucuo. The hydrogen-bonding interactions possible with solvent eliminate nonnative interactions found in uucuo, presumably through offering alternative and solvated native protein-like bonding arrangements. Levitt and Sharon (1988) divided the solvent into four classes: I, water less than 3.2 hi from a polar protein atom; 11, water less than 4.5A from a nonpolar protein atom; 111, water not in classes I, 11, or IV; IV, water bounded by the sides of the rectangular box and an ellipsoid that just fits into the box. The 107 water molecules of class I showed an average total energy that was 1.9 kcal/mol below that of bulk water. Class I waters have unfavorable interactions with other waters and slightly strained geometries, but these effects are more than offset by a strongly favorable water-protein interaction. The class I1 waters, corresponding to those contacting nonpolar elements of the surface, show a smaller relative

114

JOHN A. RUPLEY AND GIORGIO CARER1

X

X

x

X

x x x x x x . x x = x x

xx

X

I

I

X

X

8

xxx

xx

8..

x

X

X

X

m

X

L.U. V.l

-

m x

I xx X

His

x

X

x

X

X

x

x

x X

x

X I

(ns)

X X

x

I I . I I P

Glu ASPGin. Asn

.

Cvr. Thr su.

m

X

8

m I x= xx

x

8 X

x xxx

X

a x

x x I .

x(1111x

Ik. Val.

x

= I

ph..

Leu.

X

X

I x

=a Ix

x

x

x

.x x.

x 8

x

sj (ns)

FIG.35. Effect of water on parvalbumin dynamics. Time constants ( 7 ; ) were determined from time correlation functions for the vector between the two outermost nonhydrogen atoms in each side chain, ordered by residue type. (Top) In vucuo simulation of parvalbumin; (bottom) simulation with waters. From Ahlstroem et al. (1987).

binding energy (0.3 kcal/mol). Class I and class I1 waters show rates of diffusion 1.5 to 4 times less than the class IV (bulk) water value. T h e residence time for a water within a class is 4 psec for class I and 1 psec for class 11. This high mobility is countered by a high probability of return after leaving a class: 34% of the original class I water molecules were in class I at the end of the 210-psec simulation. Estimates of the entropies for class I and class I1 waters correspond to

PROTEIN HYDRATION AND FUNCTION

115

contributions to the free energy of 0.8 and 0.4 kcal/mol, respectively. These entropies, combined with estimates of the energies, give Gibbs free energies of - 1.1 and + 0.1 kcal/mol for transfer from the bulk solvent to surface classes I and 11. The experimental value for the transfer free energy (Section VI) is -0.5 kcal/mol, averaged over all of the hydration shell, probably equivalent to the sum of class I and class I1 waters. Analysis of the packing of the solvent showed that the water clustered close to the surface, giving a high density of 1.25 g/cmg for about 150 waters. This clustering was in the direction perpendicular to the surface. The waters were not brought close together in a direction parallel to the surface. T h e experimental value for the density of the hydration shell (Section VI) is 1.1 g/cm3. BPTI has been a favored subject for simulations. Earlier work with BPTI-solvent systems has been cited by Levitt and Sharon (1988).

B . Monte Carlo Simulations T h e aim of Monte Carlo simulations is to arrive at a statistical average for properties of interest by using an algorithm that appropriately samples the equilibrium ensemble. The time evolution of the system, described by molecular dynamics, is not defined by Monte Carlo simulations, but in compensation the computation is simpler and more certain of describing an equilibrium state. Monte Carlo and molecular dynamics simulations use similar potential functions. Hagler and Moult (1978) applied the Monte Carlo method to the unit ceH of the triclinic lysozyme crystal, consisting of 303 water molecules, three ordered nitrate ions, and 1001 nonhydrogen protein atoms. Of the 80 ordered waters that had been located in the crystal structure analysis, 49 were identified in the simulation. Water adjacent to the protein was significantly perturbed in its energy. The energy distribution had lowand high-energy tails, which were not found for bulk water (Fig. 36). Hermans and Vacate110 (1980) modeled BPTI in the crystal with 140 water molecules. An energy map calculated for the solvent region near the protein surface satisfactorily accommodated the 47 waters located by diffraction analysis. Only four of the crystallographic waters were not located in the simulation. The solvent was highly structured. T h e density map generated from the simulation had 158 maxima, more than the number of solvent waters. The distribution of the energies of the water molecules and their interactions is shown in Fig. 37. T h e biphasic distribution for the water-protein interaction energy is expected for a surface comprising both hydrogen-bonding and nonpolar regions.

f

2.0-

a

i c

FIG. 36. Energy distributions of waters in different environments in the triclinic lysozyme unit cell. (a) All waters, total energy (-) and energy partitioned into waterwater (*-.*) and water-protein (---) contribution. (b) Waters in close contact with the protein (. . .) and water in bulk water (-). (c) Water far from the protein (----)and water in bulk water (-). From Hagler and Moult (1978).

PROTEIN HYDRATION A N D FUNCTION

117

ENERGY (kcal/mol)

FIG. 37. Distribution of energies of water molecules about bovine pancreatic trypsin inhibitor in the crystal. Total energy (0)partitioned into contribution from water-water (0) and water-protein (0) interactions. From Hermans and Vacatello (1980).

C . Accessible Su$me and Thermodynamics of Hydration Lee and Richards (197 1) defined the solvent-accessible surface as that described by the center of a solvent molecule rolled about the surface of a solute molecule. This concept and its offspring are the basis of various computations of the contribution of hydration to the thermodynamics of protein folding and the thermodynamics of protein interactions. Edsall and McKenzie (1983) reviewed the literature on the thermodynamics of transfer processes used to model hydration and correlated this information with protein structure results analyzed according to the Lee and Richards formalism. Other notable reviews are those by Richards (1977), Nemethy et al. (1981), and Nemethy (1986). There have been several refinements in the Lee and Richards algorithm and its use. Wodak and Janin (1980) developed an approximate function, and Richmond (1984) developed an exact analytical function for the solvent-accessible surface. Both functions can be coded for machine computation. Richmond (1984) drew attention to the relationship between accessible surface and excluded volume. The simplest use of an estimate of the solvent-accessible surface is the specification of a residue as exposed or buried, depending on whether

118

JOHN A. RUPLEY AND GIORGIO CARER1

its accessible surface exceeds some threshold value. Categorizations of this type are the basis of hydrophobicity scales for amino acid side chains. Such scales give the probability of exposure for each amino acid residue, calculated as an average from accessible surface analyses of the crystal structures of a set of proteins. The binary classification of a particular residue as buried or exposed has been refined by Guy (1985) and by others (Edsall and McKenzie, 1983) through use of finer-grained categories that describe the extent to which a residue is buried or exposed. Guy (1985) used a six-layer model, defined by a set of concentric ellipsoids (Prabhakaran and Ponnuswamy, 1980). The concentration of a residue as a function of distance from the center was fit to an equation with four parameters, one of which is an apparent transfer free energy (i.e., a free energy for transfer of a residue from maximally exposed to maximally buried). Estimates of the latter parameter agreed well with experimental values of the free energy of transfer to octanol. Radzicka and Wolfenden ( 1988) measured transfer free energies from cyclohexane to water for the common amino acid side chains. Previous measurements (Wolfenden et al., 1981) had evaluated the transfer free energies from the vapor phase to water. Combination of the two sets of data evaluated the transfer from vapor phase to cyclohexane. The latter process should model the establishment of pure van der Waals interactions, and the generally negative free energies of transfer thus should be proportional to the surface areas of the side chains, which was found to be true. The transfer from cyclohexane to water was a good model for the distribution between the interior and exterior of a protein, judged by the familiar correlation with the average solvent accessibility for each type of residue. Atomic and group additivity schemes, derived from solvent accessibility calculations and measurements on model systems, have been used to estimate the thermodynamics of hydration of proteins and peptides (Eisenberg and McLachlan, 1986; Ooi and Oobatake, 1988a; Ooi et al., 1987).

Eisenberg and McLachlan ( 1986) constructed a hydration function with five atomic solvation parameters, which correspond to atomic transfer free energies. The parameters are evaluated by merging experimental data for the transfer of amino acid residue analogs from octanol to water with estimates of the solvent accessibility of peptides. The fiveparameter model satisfactorily fits the data. Application of the parametrized function to two proteins, an immunoglobulin fragment and hemerythrin, gave estimates of the solvation contribution to the free energy of folding, from an extended conformation to the native conformation, of about - 100 kcal/mol. A calculation for folding to a strongly incorrect conformation-namely, grafting the side chains of one protein onto the

PROTEIN HYDRATION AND FUNCTION

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backbone of the other, with the backbone conformation kept constantgave 17-34 kcal/mol less favorable solvation contributions. Nemethy and Scheraga and colleagues (Kang et al., 1988)worked with the hydration shell model as a means of introducing solvent into conformational energy calculations (see Kang et al., 1988, and references cited therein). The additivity function gives the free energy of transfer from vapor phase to aqueous solution. A variant of this method, with weighting by solvent-accessible surface instead of accessible volume, has been applied to hydration of proteins and to calculation of the hydration contribution to protein unfolding (Ooi et al., 1987). For ribonuclease A the free energy of hydration of the native protein was calculated as -233 kcal/mol, and that of a single extended conformation as - 535 kcal/mol (the contribution from chain statistics was not included). This treatment has been extended to estimate directly the changes in free energy, enthalpy, and heat capacity of unfolding (Ooi and Oobatake, 1988a,b). The hydration contribution to unfolding was closely linear in the number of atoms in a protein for all three thermodynamic functions. The same formalism has been applied to protein-protein and protein-peptide interactions, such as the insulin, hemoglobin, and chymotrypsin dimers (Ooi and Oobatake, 1988~). Values of the hydration free energy for each type of amino acid residue have been calculated as averages over a set of 113 proteins (Oobatake and Ooi, 1988). The results of Ooi, Nemethy, and Scheraga are for the hydration of an in uacuo molecule, and thus their values should be comparable to the experimental measurements of protein hydration from the dry powder. The calculated values for the free energy of hydration appear to be about 50% larger than the experimental. For ribonuclease the calculated value of -233 kcal/mol can be compared with the estimate from the sorption isotherm of proteins: -0.5 kcal/mol of water multiplied by about 300 mol of water per mole of ribonuclease. Because additivity calculations appear to be more accurate for higher derivatives of the free energy, comparison of calculation and experiment for the heat capacity and enthalpy of hydration of the native protein would be of interest. The solvation free energy calculated by Eisenberg and McLachlan (1986) is a transfer value that includes complex changes in interactions of side chains with solvent and with other side chains. It is not apparent how their calculated values can be compared with experimental values for protein hydration obtained outside this framework of transfer experiments. Ben-Naim et al. (1989a) provided a theoretical framework for separating the solvation thermodynamics into their several components: (1) hard-core interactions, which depend on the volume of the solute and the cost of making a cavity in the solvent; and (2) interactions of the

120

JOHN A. RUPLEY AND GIORGIO CARER1

solute surface with solvent, which can be divided into several types (van der Waals, hydrogen bond, and charge-charge). They found that surface hydrogen-bonding interactions dominate the hydration free energy of proteins. An assumption in this analysis is that the surface groups are independently solvated. Ben-Naim et al. (1989b) concluded, from an analysis of the distances between surface groups of a set of proteins, that this is true.

D . Other Simulations 1 . Ab Znitio

Clementi (1985) described ab in& computational chemistry as a global approach to simulations of complex chemical systems, derived directly from theory without recourse to empirical parametrizations. The intent is to break the computation into steps: quantum mechanical computations for the elements of the system, construction of two-body potentials for the interactions between them, statistical mechanical simulations using the above potentials, and, finally, the treatment of higher levels of chemical complexity (e.g., dissipative behavior). This program has been followed for analysis of the hydration of DNA. Early work by Clementi et al. (1977) established intermolecular potentials for the interaction of lysozyme with water, given as maps of the energy of interaction of solvent water with the lysozyme surface.

2 . Free-Energy Simulations One may wish to know the free-energy difference between closely related thermodynamic states (e.g., the effect of mutation in a protein on the binding of substrate). Separate computations of the energies of the several states and comparison of the results give a difference value with large error, as expected for a small difference between large numbers. Alternatively, one may obtain the difference value directly, through conversion of one state into the other along some continuous path, which may, but need not, have physical significance. The theoretical background and the methods for carrying out the latter process have been described by Mezei and Beveridge (1986). One approach is to define a “coupling parameter,” constructed so that the energy as a function of the coupling parameter changes smoothly, from that for the first state to that for the second, as the value of the coupling parameter changes from 0 to 1. In practice, a computationally plausible path is constructed between the two states, and it is sampled by molecular dynamics or Monte Carlo simulations at intervals sufficient to describe a smooth transition between initial and end states. Over each interval the states differ

PROTEIN HYDRATION AND FUNCTION

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by only a small perturbation, and the difference in free energy is a simple function of the difference in energy. Integration over the full path gives the total difference in free energy. A principal point is that computations with solvent explicitly included are tractable. Examples of the use of these methods have been cited by Mezei and Beveridge (1986). Several other examples follow. Bash et al. (1987) applied the thermodynamic perturbation method to complexes of thermolysin with a phosphonamidate [Cbz-GlyP-(NH)Leu-Leu] and the corresponding phosphonate inhibitor [Cbz-GlyP-(0)Leu-Leu]. T h e perturbation was carried out by using 20 windows, with 2-psec molecular dynamics simulations in each window. Computations were for the ligand in solution and bound to the enzyme. T h e solvation of the enzyme was represented by a spherical cap of 168 water molecules about the bound inhibitor. The difference in free energy of binding of the two inhibitors was calculated to be 4.38 kcal/mol, to be compared with the experimental value, 4.10 kcal/mol. These calculations point out the importance of solvation effects, which are seen in the 3.4 kcal/mol difference between the N H and 0 forms of the inhibitor. Kollman et al. (1987) summarized similar analyses, comparing transition states and preequilibrium complexes of native and mutant species of subtilisin, trypsin, and triose-phosphate isomerase. Gao et al. (1989) simulated the effect of a mutation in the hemoglobin interface, Asp Gl(99)P +. Ala, on the free energy of cooperativity in binding oxygen. They calculated a net effect of - 5.5 kcal/mol of interface, to be compared with an experimental value of - 3.4 kcal/mol. T h e relatively small net effect was the result of compensating large contributions of varied sign. The free energy change associated with the effect of the mutation on solvation was 46 kcal/mol for the deoxy species and 68.5 kcal/mol for the oxy species. The solvation effects were the largest. The computations revealed the substantial thermodynamic contributions of individual molecular level events, hidden beneath a relatively small net effect of the mutation. Warshel and collaborators (Warshel and Sussman, 1986; Warshel et al., 1988) developed the empirical valence bond method for obtaining freeenergy differences and activation free energies. The effects of Gly-toAla mutations in trypsin were accurately simulated. This method was earlier applied to calculation of the potential surface for general acid catalysis of a disaccharide in solution and bound to lysozyme (Warshel and Weiss, 1980). Warshel et al. (1986) calculated protein pK values in solution by using a microscopic model and a reversible charging process. Jorgensen (1989) carried out Monte Carlo simulations of the Nmethylacetamide-chloroform and N-methylacetamide-water systems.

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JOHN A. RUPLEY AND GIORGIO CARER1

Thermodynamic association constants for the two solvents, calculated from the potentials of mean force for association of two N-methylacetamide molecules, agreed closely with experiments. An important point is that, in water, the amide dimers are not hydrogen bonded, but rather are stacked with favorable dipole alignment so as to minimize the loss of hydrogen bonding with water. In chloroform the dimers are hydrogen bonded. This unexpected picture of the dimer in water perhaps explains difficulties that have been found in applying the N-methylacetamide model to protein processes. 3 . Electrostatics Electrostatic effects have long been recognized as important for protein properties and function. Water strongly affects electrostatic interactions, and, conversely, it must be equally affected by the electrostatic field of the protein. Considerable attention has been paid to computation of electrostaticcontributions for proteins and other macromolecules (Gilson and Honig, 1988a,b; Honig et al., 1986; Maroncelli et al., 1989; Matthew, 1985; Warshel and Russell, 1984; Warshel et al., 1984). Generally, the solvent is modeled as a continuum or periodic structure. Churg and Warshel (1986) explicitly considered solvent-charge interactions in an analysis of the redox potential of cytochrome c. Wendoloski and Matthew (1989) used molecular dynamics simulations to generate a set of conformers of tuna cytochrome c, upon which electrostatic calculations were carried out separately. They found fluctuations of 0.32.0 pK units for individual ionizable groups. Fluctuations of this kind may be important for catalysis. It may be that detailed electrostatic calculations carried out for a dry or partially hydrated protein, for which the complexity of the solvent is minimal, would afford useful tests of the models, through comparison with experiments. VI. PICTURE OF PROTEIN HYDRATION

This section summarizes and coordinates the results of measurements described in Sections II-V. The discussion is based on a picture of the hydration process developed by Careri et al. (1980) and by Rupley et al. (1983). Others have presented similar pictures (Finney and Poole, 1984; Finney et al., 1982).Literature citations are minimal, and it is hoped that the reader will consult previous sections for more complete documentation. Table VII summarizes the chemistry of the protein at various hydration levels and also the principal points of the discussion that follows.

TABLE VII Chemistry of the Hydration Process” Hydration level ( h )

Thermodynamics

Structure

Dynamics

0-0.07 (0-60)

“Knee” in sorption isotherm Large differences in partial molar quantities for transfer of water from bulk solvent to hydration layer; average values: ACl = - 1.5, ABl = -4 kcal/mol Heat capacity of adsorbed water between that of ice and liquid: Zp, = 0.8 cal K-Ig-’ Folded state stable; T, solution value At low hydration protons redistribute among ionizable groups, to reduce number of charges At 0.05 h: normalization of pK At 0.07 h: phase transition (2-dimensional condensation)

Structure of protein at low hydration same as that for full hydration, at 1 A level of comparison Domain displacements of 1-2 A at low hydration Computer simulations show general contraction and some H-bond rearrangement in uacuo Water interacts principally with charged groups (ca 2 watedgroup) At 0.07 h : transition in surface water, from disordered to ordered and/or from dispersed to clustered state; detected in IR, ESR, neutron scattering, thermodynamic, and other measurements; associated with completion of charged group hydration

Water mobility low Mobility of noncovalently bound ligand constant and essentially frozen from 0 to 0.2 h ; T = 4 x 10-9 sec Protein motions also frozen Enzymatic activity negligible

0.07-0.25 (60-220)

Plateau in sorption isotherm Small differences in partial molar quantities for transfer of water into hydration layer, which, with increasing hydration, decrease to near 0; average values: ACl = -0.2, AR, = - 1 . 1 kcal/mol, AFl = 0 for h 2 0.15

Water interacts principally with polar protein surface groups (ca 1 water/ polar site) Water clusters centered on charged and polar sites Clusters fluctuate in size andlor arrangement

Internal protein motion (H exchange) increases from 1/1000 at 0.04 h to full solution rate at 0.15 h At 0.1-0.15 h: chymotrypsin and some other enzymes develop activity At 0.15 h : long-range proton movements along percolative networks, seen in dielectric measurements (continued)

TABLE VII (continued)

Hydration level (h)

w

E3 rp

Thermodynamics

Structure

Dynamics

Heat capacity of adsorbed water is greater than for bulk solvent: Fpl = 1.4 cal K-’g-’ T,,, decreases strongly with increased hydration

At 0.15 h: long-range connectivity of the surface water is established, in 2-dimensional percolative phase transition. Network of H-bonded water spans protein surface; the network has fluctuating and random connectivity, richness of connections increasing with hydration level

0.25-0.38 (220-305)

Region of strong upswing in sorption isotherm, corresponding to thermodynamic quantities being close to bulk solvent values Phase transition (condensation) At low temperature ice crystals form at h 2 0.34 (nonfreezing water)

At 0.25 h: start of condensation of water onto weakly interacting unfilled patches of protein surface; seen in dynamic and thermodynamic properties

For lysozyme parallel increase in enzyme activity and motion of noncovalently bound ligand Water motion increases strongly with increased hydration

0.38 (305)

Full hydration, defined as point at which major changes in thermodynamic properties are complete Table V I l l gives average thermodynamic differences between hydration and bulk water, which are 10- 15% of bulk water values and less than ambient thermal energy for free energy T,,, is close to solution value

Monolayer of water covers surface Interaction with charged and polar surface groups selects locally ordered arrangements of hydration water Fluctuation between various instantaneous arrangements, as in liquid water Arrangements mesh with the bulk solvent Arrangements cover large area (20 A*) per water molecule

Part, and perhaps most, of hydration water has mobility close to bulk water. Rotational motion is 1/10-11100 that of bulk solvent, and diffusion constant, 1/5. Few, if any, hydration water has ~~~~h~~ > sec. Considering mobility of hydration water and its small free-energy difference from bulk solvent, term “bound water” seems inappropriate

Diffraction studies show, for most proteins, clusters or threads of H-bonded water; for some proteins, extensive H-bonded networks

>0.38 (>305)

Hydration forces reflect cooperative interactions of water in several layers about macromolecule Water in layers outside monolayer adjacent to protein is perturbed an order of magnitude less strongly than water in monolayer proper: lAGll < 0.05 kcal/mol Small decrease in T, between 0.38 and 0.8 h likely represents hydration of unfolded state

sec Mobile bound ligand; T = 7 X Enzymatic activity of lysozyme 1/10 solution value Full internal motions of protein Dynamic and thermodynamic coupling between hydration water and protein, seen in Mossbauer spectroscopy and computer simulations Some motional properties change significantly above hydration level for completion of principal changes in thermodynamic properties: motion of bound ligand (ESR); enzyme activity; water motions (NMR) Motion of large groups at protein surface may require participation of adjacent bulk solvent (e.g., in cooperative H-bond rearrangements)

"Values in the first column give a range of hydration (in grams of water per gram of protein and, in parentheses, moles of water per mole of lysozyme). Data and descriptions are mostly from experiments on lysozyme, cited in the text. Thermodynamic values with subscript 1 are for the water component. (Adapted from Table I1 of Rupley et al., 1983.)

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A . Fully Hydrated Protein 1 . Solvent

Table VIII gives the thermodynamics of transfer of water from the bulk solvent into the protein-water interface. The values were estimated from selected measurements on three proteins-lysozyme, ovalbumin, and ribonuclease-which are likely to be typical. The sorption isotherms, and hence the free energies of hydration, are similar for globular proteins (Kuntz and Kauzmann, 1974). The partial specific heat capacities of proteins are also similar (Suurkuusk, 1974; Yang and Rupley, 1979). There is only one measurement of the hydration dependence of the specific volume (Bull and Breese, 196813). There is also only one calorimetric measurement of the hydration dependence of the heat of solution (Almog and Schrier, 1978), which is judged to give a more solid value of the heat of transfer than van’t Hoff analyses of the temperature dependence of sorption isotherms. The values in Table VIII are averages over all water of the interface, and they express all nonideality of the system, regardless of whether at the microscopic level the nonideality might be seen to contain a substantial contribution assignable to a protein process, such as redistribution of protons among ionizable groups. The thermodynamic properties of the hydration shell (Table VIII) show it to be slightly, but not strongly, different from the bulk water. The free-energy difference is only 0.5 kcal/mol of water, slightly less than the ambient thermal energy. The heat capacity, enthalpy, and volume changes associated with hydration are 10-15% of the bulk water values. About 300 water molecules are sufficient to cover the lysozyme surface. This is a remarkably small amount of water. Calculations based c p the crystal structure of lysozyme show that the surface is about 6000 A2 in area (Lee and Richards, 1971; Shrake and Rupley, 1973). Thus, each water covers, on average, about 20 A 2 , which is twice the “projection” of a water molecule packed as in the liquid. Since 20 Azis the most area a water molecule can cover and maintain hydrogen bonding, there can be no multilayer water. Moreover, whatever arrangements there are at the surface must integrate simply into the bulk water; namely, there is no “B” shell of disordered water, required to interface the water adjacent to the protein surface with the bulk solvent. It appears possible that the protein surface selects local arrangements from among the many possible in bulk water, so satisfying at once the requirement of large area covered per hydration water and also the requirement of meshing with the surrounding solvent. This suggestion is consistent with the considerations of the preceding paragraph and with

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TABLE VIII Thermodynamics of Transfer of Solvent into Interface a

Thermodynamic variable Free energy Enthalpy Entropy Volume Heat capacity

Difference (hydrated protein - pure components)

- 0.5 kcal/mol* - 1.4 kcal/mol -3.0 cal K-' mol-' - 1.7 ml/mole 2.5 cal K-I mol-' f

"The difference values are the change in thermodynamic variable for the hydration process, that is, the addition to dry protein of the amount of water required to reach the point of full hydration. The values are calculated per mol of water. This formalism attributes all nonideality to the solvent. *Calculated by integration of the sorption isotherm measured for lysozyme at 27°C (Hnojewyj and Reyerson, 1959). 'From a calorimetric measurement of the heat of solution of lyophilized ribonuclease at pH 7.0 and 25°C (Almog and Schrier, 1978). dFrom the difference between the free energy and enthalpy of hydration. The two small globular proteins, lysozyme and ribonuclease, perhaps can be considered compatible at this level of comparison. CFromdensity measurements on ovalbumin at 25°C (Bull and Breese, 1968b). /From calorimetric measurements of lysozyme at 25°C (Yang and Rupley, 1979).

the following observations: ( 1) infrared spectroscopic and thermodynamic measurements suggest that the polar and charged groups are the primary sites of hydration and are filled at hydration levels below 0.25 h. (2) The average spacing of polar and charged groups on a protein surface is 4-5 A. (3) Layers of hydrogen-bonded water present in ice I, orthogonal to the c axis, show a projected area of 20 hi2, with 4.5 A separation between water molecules. The crystallographic results of Blake et al. (1983), who analyzed highresolution X-ray diffraction data for human lysozyme (HL) and tortoise egg white lysozyme (TEWL) fit the above picture. The crystallographic

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estimate of surface water is in good agreement with the 300 molecules of water seen by the heat capacity measurements. The area covered per water molecule is estimated as 19-21 Hi2. The ordered water is in a monolayer about the protein surface. The average number of hydrogenbonded neighbors is two or three. If the interaction between the ordered water in the monolayer and the bulk solvent is taken into account, then, on average, the number of hydrogen-bonded neighbors for water in the surface monolayer is as for bulk water. The waters are nearly all bonded to polar or charged protein atoms. In general, charged side chains bind two ordered waters; polar groups, one. The observation of localized water in X-ray diffraction studies does not mean that the water is strongly bound. Indeed, the hydration water is weakly bound, both with regard to the equilibrium thermodynamics of transfer from the bulk solvent, for which the free energy is slightly less than the ambient thermal energy, and with regard to residence time on the protein, shown by NMR measurements to be lo-* sec or shorter. Measurements of the dynamic properties of the surface water, particularly NMR measurements, have shown that the characteristic time of the water motion is slower than the bulk water value by a factor of less than 100. The motion is anisotropic. There is little or no irrotationally bound water. Study of a protein labeled covalently with a nitroxide spin probe (Polnaszek and Bryant, 1984a,b) has shown that the diffusion constant of the surface water is about 5-fold below the bulk water value. The NMR results are in agreement with measurements of dielectric relaxation of water in protein powders (Harvey and Hoekstra, 1972). Networks formed of hydrogen-bonded water and protein surface atoms have been described by X-ray diffraction studies. Various other measurements (e.g., heat capacity and neutron scattering) have given evidence of water clusters. Protonic conduction over the surface of lysozyme, displayed in dielectric measurements, appears to be associated with statistical paths and long-range connectivity characteristic of fluctuating network arrangements of the surface water. There are preferred paths of proton movement, even for the fully hydrated protein, that lead, for lysozyme, to a disproportionately large flux of protons through the active site. Although protonic conduction may be of interest in its own right, it can properly be viewed as a probe of the long-range connectivity of the hydrogen-bonded surface network, and equally important, of the percolative character of the network. The general physical model of percolation theory applies to a broad range of phenomena that are rooted in spatially random events and topological disorder and that reflect long-range connectivity. One can picture the protonic process as being a transfer along threads of hydro-

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gen-bonded water molecules adsorbed on the protein surface. At the percolation threshold there is a sharp change in the characteristic length that describes the connectivity of the surface water. Above the threshold the mean free path of the protons is long, and the threads act as shorts bypassing the local details of the protein surface. The stochastic basis of the percolation model is consistent with fluctuating paths within the water network. To summarize, the protein surface is covered by a monolayer of water. It differs only slightly in thermodynamic properties from bulk water. It is organized in local arrangements that are determined principally by the polar protein surface groups. These local arrangements appear to be a subset of those found in bulk water. Like those in bulk water, they fluctuate, but have a defined time-average structure. The surface arrangements may display special properties, such as a high probability of proton movement through the active site. Threads, clusters, or networks of water appear to be a characteristic of the surface. The motions of the surface water are one to two orders of magnitude slower than for bulk water. The picture given above is a first-order description. Clearly, water outside the monolayer participates in some protein processes. Rate processes involving large ligands or protein side chains may sense multilayer water. The change in TEMPONE motion with hydration of lysozyme is largely, but not completely, saturated at 0.38 h. Long-range hydration forces have been described for assemblies of proteins, nucleic acids, and membranes (Israelachvili and Marra, 1986; Parsegian et al., 1986; Prouty et al., 1985; Rand et al., 1985). These are cooperative phenomena, for which the perturbation per water molecule is small. 2. Protein

Contributions specifically from the protein to the thermodynamics of hydration are included in the values given above for transfer of solvent into the interface. Protein rate processes are strongly affected by hydration. The dry protein shows greatly reduced internal motions, measured by Mossbauer spectroscopy, neutron scattering, fluorescence spectroscopy, and other methods. Surface motions, monitored by spin probes or spin or Mossbauer labels, are similarly frozen in the dry protein. The following paragraphs comment on the appearance of motion characteristic of the hydrated protein and on the coupling between protein and solvent motions. Above 0.2-0.25 h the rotational motion of the spin probe TEMPONE grows explosively, with an apparent 15th-order dependence on water

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activity. These measurements suggest that the development of surface motion is tied to the condensation event that completes the hydration shell, seen in the heat capacity isotherm as the 0.25 h discontinuity. This correspondence appears to be physically plausible. For motion of a relatively large molecule at the protein surface, and presumably for motion of protein side chains at the surface, many interactions with the protein must be broken and reformed. The presence of a substantially continuous water region should facilitate this process, through the intermediate formation of protein-water and ligand-water interactions. Several observations demonstrate the coupling of internal motions of the protein with the adjacent solvent. (1) Protein motions detected by Mossbauer spectroscopy, understood to be of helical segments or other blocks of atoms, are restricted in the absence of water. The glass transition at 180-200 K is not observed, or the changes associated with it are small for the dry protein (Krupyanskii et al., 1982; Parak, 1986). (2) Neutron scattering data are consistent with the Mossbauer measurements (Cusack, 1989). (3) Changes with temperature in the internal motions of myoglobin are paralleled by changes in solvent motions detected by Mossbauer spectroscopy, high-frequency dielectric measurements, IR spectroscopy and heat capacity (Doster et al., 1986; Parak, 1986), and ESR spin probe motions (Steinhoff et al., 1989). (4) Solution measurements of various processes (e.g., enzyme activity and the interaction of oxygen with heme proteins) have shown that the reaction rate is reduced at high solvent viscosity (Beece et al., 1980; Gavish and Werber, 1979). (5) Molecular dynamics simulations, comparing proteins in aqueous solution and in vacuo, have indicated that solvent produces about a 10-fold reduction in the time constants for motions of side chains (Ahlstroem et al., 1987) and a 2-fold reduction for all atoms (Levitt and Sharon, 1988). (6) The mechanical properties of lysozyme crystals are linked to the sorption equilibria. Several elementary processes have been suggested as being important for coupling protein and solvent motions. (1) Groups of protein atoms or domains of the protein undergo restricted Brownian motion, for which solvent contributes to the frictional dissipative processes and also is the source of the random fluctuating force driving the motion (for a review discussion see Gavish, 1986). (2) The fluctuating electric field of the hydration water, arising from water dipole reorientation, might interact with the fluctuating electric field of the protein charged groups (Singh et al., 1981) or with the charge distribution on the amide backbone (Careri et al., 1979a), which could cross-correlate, in turn, with transitions between conformational substates. (3) The surface of a protein has networks of hydrogen-bonded protein and solvent atoms. Coop-

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erativity within these clusters might provide the coupling between conformational motion and water fluctuations (Doster et al., 1986). (4) T h e elementary processes occurring at the protein surface and having time constants in the nanosecond range, which are those likely to couple with motions of protein groups or domains, have been listed as the following (Careri, 1974; Careri and Gratton, 1986): proton transfer reactions involving the hydration water, charge fluctuations of the ionic medium, side-chain motions, and motions of hydrogen-bonded surface networks. In the absence of water, the nanosecond and slower motions of the protein are essentially frozen, as they are at temperatures below 180 K. T h e protein is likely in a glassy state, in which large-scale collective motions are inhibited and the allowed dynamics are picosecond and subpicosecond atom motions. Scanning calorimetry shows that, as the hydration water is removed, the transition temperature for denaturation rises. At low hydration levels unfolding is not detected at accessible temperatures. Large-scale motions such as tryptophan ring rotation are not detected in the dry or nearly dry protein (Careri and Gratton, 1986). A small amount of water acts as a catalyst, facilitating conformational transitions by participating in alternative hydrogen-bonding arrangements, so lowering the energy barrier for the conformational transitions (Chirgadze and Ovsepyan, 1972a). This role of water as a catalyst, in which it functions as a reactant in hydrogen bond exchange processes, is distinct from its role in the fluctuation processes discussed in the preceding paragraph. B . Hydration Process 1 . Time-Average Properties

The process of protein hydration is the stepwise addition of water to dry protein, until the hydration end point is reached. Heat capacity measurements (Yang and Rupley, 1979) serve as a framework on which to develop a picture. Figure 38 gives the dependence on hydration level of various time-average properties of lysozyme, over the hydration range 0-0.4 h, from the dry protein to slightly beyond the end point of the process. Curve d shows the dependence of the apparent specific heat on hydration level. It is directly related to the extent to which the thermal response of the lysozyme-water system deviates from ideal behavior. T h e nonideality of the system shows three discontinuities: at 0.07, 0.25, and 0.38 h. Below 0.07 h thermodynamic and IR spectroscopic measurements have shown that water adsorbed to the protein is localized at charged

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JOHN A. RUPLEY AND GIORGIO CARER1

I'

C

I

0

0.I

0.2

f ' I

I

I

I

I

I

0.3

0.4

gH20/g PROTEIN

FIG.38. Comparison of heat capacity and spectroscopic properties. Effect of hydration on lysozyme time-average properties. (Curve a) Carboxylate absorbance (1580 cm - 1 ) ; (curve b) amide I shift (-1660 cm-I); (curve c) OD stretching frequency (-2570 cm-I); (curve d) apparent specific heat capacity; (curve e) diamagnetic susceptibility.From Rupley et 01. ( 1983).

sites. Nucleation theory (Abraham, 1974) states that it is impossible to condense water on an insoluble particle with a radius less than 100 A, unless the pressure is greater than the equilibrium vapor pressure of water. Only because soluble elements such as ionizable residues are present on the protein surface can water vapor condense on the macromolecule at low water activity to initiate the hydration process. At the 0.07 h discontinuity, the heat capacity function shifts from generally downward-trending to strongly upward-trending. This is expected for a two-dimensional condensation process-here, the formation of mobile water clusters from dispersed water associated with ionizable protein surface groups. This transition in the surface water is seen also in the IR spectroscopic properties (Fig. 38) and other proper-

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ties (Middendorf et al., 1984; Rupley et al., 1980). Between 0.07 and 0.25 h the water clusters grow, and there is no qualitative change in the chemistry of the surface. At the 0.25 h discontinuity, condensation of water over the weakest interacting regions of the surface begins and proceeds until completion of the hydration shell at 0.38 h. This event is a typical condensation. It occurs within a narrow range of water activity, near unity, and is associated with addition to the surface of about one-third of the total hydration shell. The 0.38 h discontinuity corresponds to the hydration end point, above which the nonideality per mol of protein is constant. The heat capacity measurements define the hydration end point tightly. These measurements are sensitive to the interactions of water with both hydrogen-bonding and nonpolar protein surface’groups and should reflect essentially all time-average chemistry associated with hydration. The rise and fall of the apparent heat capacity between 0 and 0.07 h is a transition heat contribution, associated with the transfer of protons from carboxylic acid to basic protein groups. As the protein is dried to low hydration levels, below about 0.05 h, the system reacts to a high net charge in the unfavorable low-dielectric environment of a protein surface stripped of water. Charge is neutralized by changing the effective pK order, so carboxylate groups become among the most basic of the ionizable protein groups. Infrared measurements show this transition in the carboxylate pK (Fig. 38). Implicit in the above description is an evolution, during the hydration process, of the properties of the protein and especially of the surface water. The water environment changes from isolated adsorbate, on charged sites, to mobile clusters, to complete monolayer. The chemistry of the water also changes (see the IR and heat capacity measurements of Fig. 38). One should not expect the thermodynamics for vaporization of a single water molecule bound at low hydration to a charged group to be the same as the thermodynamics for removal of one member of a shell of water about that same charged group in the fully hydrated molecule. There should be very little “tightly baund” water at full hydration, certainly not the 0.05-0.1 g / g adsorbed below the knee of the sorption isotherm. It is also not correct to divide the hydration water into types, based on the steps of the hydration process. The division of hydration water into A and B shells, by analogy with ion hydration, also appears to be incorrect. The A shell, the interface adjacent to the protein, is arranged so that it integrates smoothly with the surrounding solvent, and thus there is no B shell of water with properties that differ from the bulk water by as much as the ambient thermal energy.

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Time-average properties develop with hydration in accord with the above three-stage picture. The knee of the sorption isotherm corresponds to the low hydration condensation event; the upswing in the sorption isotherm, to the high hydration event. The IR measurements (Fig. 38) of intensity of the carboxylate band, the amide I band, and the O-D stretching frequency all show a discontinuity at the 0.07-h transition. The IR measurements also show that the hydrogen-bonding sites are largely filled before the 0 . 2 5 4 transition, as is expected if the transition is condensation of solvent over the weakest-interacting portions of the surface. The hydration dependence of the diamagnetic susceptibility, which is similar to the dependence of the heat of transfer of water into the interface, parallels the filling of the hydrogen-bonding sites. The above picture should be general. As noted, globular proteins are closely similar in their sorption isotherms, and heat capacity measurements in the water-poor region, for several proteins, are consistent with the data of Fig. 38. This picture is in accord with theory. Hill (1949) treated the case of adsorption on a heterogeneous surface with the further restrictions that the adsorption be localized (i.e., that the energy of interaction between surface and adsorbate be large compared to the thermal energy) and that the adsorption be unimolecular. There can be lateral interactions among adsorbate molecules on the surface. This model applies in detail to a protein surface and the interaction of water with it: the strong localizing interactions with the surface and the lateral interactions are provided by water hydrogen bonds; the hydration shell is a monolayer; and the surface is heterogeneous, consisting of charged, polar, and nonpolar sites. The Hill treatment predicts two phase transitions: one at low coverage of the surface, which is a two-dimensional condensation of water dispersed about the surface into clusters; the second at high coverage, where adsorbate condenses to complete the monolayer. These are the transitions found for globular proteins.

2 . Dynamics The dependence of protein and solvent dynamics on hydration fits well into the above three-stage picture for some, but not all, properties. For dynamic properties that do not fit well, analysis on a case-by-case basis within the framework of the time-average picture can be informative. For example, consider protonic conduction, measured by the megahertz frequency dielectric response for partially hydrated powders of lysozyme. The capacitance grows explosively above a hydration level of 0.15 h, in a way characteristic of a phase transition (Section 111,A). The hydration dependence of thermodynamic properties shows, however,

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that there is no phase transition between 0.1 and 0.25 h. This apparent conflict between the dielectric results and the time-average picture was resolved by demonstrating that protonic conductivity is a percolation process. A percolative transition reflects the connectivity within the system and is independent of the chemistry of the elements. The critical hydration level, 0.15 h, is simply the point of half-coverage of the surface with water and, according to the percolation model, the point at which long-range connectivity is established. There is an impression, which is likely correct, that the hydration dependence is more complex for dynamic properties than for timeaverage properties. Several examples follow. 1. ESR measurements of noncovalently bound TEMPONE show the high-hydration event at 0.25 h as controlling motion, but the rotational correlation time continues to change above the time-average hydration end point at 0.38 h. The latter change may be due to residual restrictions on rotation of a large group within a two-dimensional surface monolayer, which would be released in a three-dimensional solution environment. An effect of this kind should be found generally for motions of groups of atoms at the protein surface (e.g., protein side chains or bound substrate). 2. Dielectric and NMR measurements of the water in powder samples also show both the 0.25 h high-hydration event and changes above the time-average hydration end point. The changes in dielectric properties above 0.38 h appear to be a function of the properties of the bulk material (Careri et ul., 1985). The changes in NMR properties (Section 111) are more interesting. Changes in correlation time above 0.38 h are perhaps most simply explained as being due to collective motions of water molecules, which would not be sensed in most thermodynamic measurements, owing to the small perturbation per water molecule. 3. Amide hydrogen exchange in protein powders depends weakly on water activity, and its hydration dependence is complete within the low-hydration region (0.15 h). Apparently, the rate-determining step for the exchange of buried hydrogens is not much influenced by the protein surface. This is unexplained. 4. There appears to be no single pattern describing the hydration dependence of enzyme activity. Lysozyme activity is correlated with the unfreezing of surface motion at 0.25 h and also with the onset of surface percolation. There are changes in activity above 0.38 h, such as the changes found for rotational motion of TEMPONE. The hydration threshold for chymotrypsin activity, at 0.12 h, is substantially lower than that for lysozyme. A correlation with percolation is an attractive, but untested, possibility.

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5. Changes in dynamic properties measured at temperatures between 200 and 273 K may not correlate well with the model discussed here, which is based on room temperature thermodynamic measurements. Perhaps this is not surprising, in view of the dependence of the dynamic transitions on both temperature and hydration level.

3. The 200 K Transition: Temperature and Hydration Dependence of Dynamics

Various properties of the hydration water and the protein show a transition near 200 K that depends on hydration level. The hydration water properties include (1) the frequency of the O-D IR band of the solvent in myoglobin crystals (Doster et al., 1986); (2) the specific heat of water in myoglobin crystals, with a glass transition near 220 K (Doster et al., 1986); (3) the dielectric relaxation time of water in myoglobin crystals (Singh et al., 1981); (4) the relaxations of water adsorbed on protein powders observed by NMR (Andrew, 1985; Bryant, 1988); (5) the relaxations of water in crambin crystals observed by NMR (Usha and Wittebort, 1989); (6) the motions of water in myoglobin crystals monitored by electron paramagnetic resonance (EPR) spin probes diffused into the crystals (Steinhoff et al., 1989) and motions for solutions of several proteins monitored by EPR spin probes and spin labels (Ruggiero et al., 1986); (7) Mossbauer spectra for [57Fe]ferricyanidediffused into the solvent of myoglobin crystals (Parak, 1986); and (8) temperature dependence of the linewidth broadening for RSMR spectra of bovine serum albumin (Goldanskii and Krupyanskii, 1989). The protein properties include (1) motions of several proteins monitored by ESR spin labels (Belonogova et al., 1978, 1979; Likhtenshtein, 1976; Steinhoff et al., 1989) and Mossbauer labels (Belonogova et al., 1979; Likhtenshtein, 1976); (2) temperature dependence of neutron scattering for myoglobin (Cusack, 1989; Doster et al., 1989); (3) Mossbauer spectra (Parak et al., 1988) and RSMR spectra (Goldanskii and Krupyanskii, 1989) of myoglobin; and (4)mechanical properties of lysozyme crystals (Morozov and Gevorkyan, 1985; Morozov et al., 1988). The measurements cover a wide range of characteristic times: 1 psec for neutron scattering, 1 nsec for NMR relaxation, and 100 nsec for Mossbauer spectroscopy; the heat capacity is a time-average quantity. The observation of similar responses for measurements of widely varied character suggests that understanding the 200-K transition is fundamental to understanding the dynamics of proteins. Several pictures (Doster et al., 1986; Goldanskii and Krupyanskii, 1989; Parak et al., 1988) of the events associated with the transition are combined in the following summary. At temperatures below the critical

-

-

-

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temperature of about 200 K, the protein molecules are frozen, as a glasslike solid, into individual conformational substates (Section VII). The motions are the vibrational motions of a harmonic solid. As the temperature is raised through the critical temperature, molecules become able to jump between conformational substates, and segmental motions contribute. The characteristic size of the segments is, for the Mossbauer time scale, about 5 di. It is apparently smaller for higher-frequency sampling (neutron scattering). Goldanskii and Krupyanskii (1989) suggested that the range of size of the cooperative fluctuating units is what allows a protein to seem metallike in compressibility measurements, for example, and fluidlike in other, fast time-scale, measurements, such as the diffusion of oxygen within the protein matrix. The solvent melts over a broad temperature range, in what appears to be a glass transition centered at about 220 K. It may be more appropriate to view the water transition as clusterwise melting, in which the size of the cooperativity units decreases with increased temperature. The cluster size also would decrease with a decrease in hydration level. There is coupling between protein and solvent motions. The evidence is summarized in Section VI,A,P. Temperature and hydration level are linked in determining the dynamics of protein and solvent. The dry protein shows, for all temperatures, only the restricted motion found below the critical temperature for hydrated samples. A fully hydrated sample shows strong temperature dependence for the dynamic properties of both protein and hydration water, for temperatures above the critical temperature. Partially hydrated samples behave complexly. Goldanskii and Krupyanskii ( 1989) gave a particularly good discussion of the linkage between the effects of temperature and hydration. C . Bound Water The term bound water and equivalent terms, such as hydration shell, solvent shell, or hydration water, have been commonly used to refer to water affected by the protein surface, particularly in solution measurements. The point of the following discussion is that the bound water can be defined sufficiently precisely and independently of the type of measurement to make both the term and the concept useful. Consider again the heat capacity data of Fig. 38. The isotherm defines the phase diagram of a protein-water system. Below 0.38 h there is a solution of water in protein-more accurately, water on the protem. At 0.38 h there is phase separation, and at higher hydration levels the hydrated protein phase is constant in composition. This end point is well

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defined. Time-average properties appear to change coherently with hydration. The crystallographic and thermodynamic measurements are in accord. Globular proteins show closely similar sorption processes. There appears to be no time-average property that indicates an hydration end point greater than about 0.4h for globular proteins. Dynamic properties sometimes, although not always, can be understood within the framework of the picture developed for the time-average properties. A principal point is that water adjacent to the protein differs substantially from second-layer and multilayer water. The thermodynamic properties are significantly perturbed in the adjacent layer (the free energy, by 0.5 kcal/ mol; heat capacity, volume, and enthalpy, by 10- 15%).The properties of second-layer water are about an order of magnitude less strongly affected, and the perturbation decays exponentially with distance from the surface (Israelachvili and Marra, 1986; Parsegian et al., 1986). With this picture the terms hydration shell and bound water are understood to mean the water at the hydration end point. With this definition several questions should be addressed. 1. Some give identifiers such as bound water or hydration shell an operational definition-namely, the water detected by a particular measurement as having been affected by the protein-and consequently the value assigned to the bound water will differ between measurements. Specifically, for some measurements the hydration dependence is complete at low hydration. For example, the heat of transfer of water into a protein powder is near zero at 0.2 h. How do observations of this kind bear on the bound water? It would be wrong, given the present level of understanding of protein hydration, to say that the hydration shell consists of only 0.2 g of water/g of protein, with a qualification, as detected by this type of measurement. Rather, the significance of this particular measurement is the light it sheds on the hydration process, through the correlation of the heat changes with filling of hydrogenbonding sites of the surface, detected in other isotherms and spectroscopic measurements. 2. Is the picture of bound water defined by time-average properties applicable to dynamic properties? In view of the solid understanding of the time-average behavior, it seems appropriate to use this picture as the basis for interpretation of dynamic properties. Clearly, there are difficulties in doing this. The characteristic times for motions of a protein and the adjacent water cover a wide range, and they are sampled differently according to the type of measurement. The understanding of proton NMR measurements for protein solutions may be particularly difficult, in view of the several mechanisms for relaxation. Collective motions of groups of water molecules are likely to enter into dynamic processes.

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Some apparent conflicts between the hydration dependence of dynamic properties and the time-average picture may be treated relatively simply, as noted above. There is additional discussion in Section 111. 3. Do some measurements detect the effects of hydration beyond the end point defined above? The answer is, clearly, yes. Electrostatic interactions may be longer range. Hydration forces have been discussed. Measurements of this kind are treated by use of a different model, which may not include a distinct water monolayer at the protein surface. The point to be made, however, is that water beyond the monolayer is not strongly perturbed and differs substantially from water adjacent to the surface. 4. Is the hydration shell tightly bound? The answer is, clearly, no. The average free energy of binding is slightly less than thermal energy. The motions of the bound water, although reduced compared with the sec. bulk water, are still fast, with characteristic times below 5. Is it useful to define bound water for other macromolecules? Perhaps, if there is an invariant conformation and surface at least over the range of high water activity (greater than perhaps 0.9). This is likely the case for some nucleic acids, but may not be so for saccharide polyelectrolytes. There are several nonstandard views of bound water. Ling and colleagues (see Ling, 1988, and other articles in the same issue) developed a polarized multilayer theory of cell water, in which all of the water within the cell is bound to protein and has equilibrium properties affected by the protein, leading, in particular, to the exclusion of sodium ion. This view is at variance with the description given here, in which only a monolayer has properties so strongly perturbed. Wiggins (1988; Wiggins and MacClement, 1987) suggested that water in hydrophobic clefts is organized as “stretched” water, with special thermodynamic properties. Although we have used the term bound water in this section, we have not used it generally elsewhere in this review. The word bound carries connotations of slow dynamics and large free energy of formation of a complex, which do not appropriately describe the water about a protein. The terms hydration shell and hydration water lack the above connotations and thus are preferred. D. Hydration and Conformation

There are several results that support the conclusion that the conformation of a protein, defined as the backbone and internal arrangements, is similar in the dry state and in solution, to a resolution of perhaps 1 A.

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TABLE IX Spin-Spin Interaction for Nitroxide Labels on Lysozyme a

Hydration level (grams of water per gram of protein)

1.8 Spinsnysozyme molecule 0.02 0.10 0.17 0.39 0.62 2.3 Spinsflysozyme molecule 0.02 0.08 0.167 0.321 0.64

Average separation dlld

(612)

2.4z, (GI

0.56 0.56 0.57 0.56 0.57

25 25 24 25 24

64.0 65.2 65.2 66.2 67.0

0.64 0.63 0.62 0.59 0.58

20 20 21 23 24

63.8 65.4 65.8 66.8 68.2

“Succinimidyl-2,2,5,5-tetramethyl-3-pyrrolin1-oxy l-3-carboxylate was reacted with lysozyme to give labeling of 2.3 or 1.8 mollmol of protein. ESR spectra were measured at - 160°C. The parameter d l l d (Likhtenshtein, 1976),a ratio of amplitudes, describes the extent of spin-spin interaction. The estimate of average distance between spins was made from the value of d l l d by use of the plot of Fig. 15 of Likhtenshtein (1976).The separation of the two outer hyperfine extrema (2A,,)is determined by the mobility of the spin probe and its environment. (From unpublished data of P.-H. Yang, G. Tollin, and J. A. Rupley, with permission.)

(1) Lysozyme randomly reacted with a spin label, at the level of two labels per molecule of protein, showed no change in average separation of the labels as the hydration was changed from dry protein to 0.17 h (Table IX), to a resolution of 1 A. (2) A jump hydrogen-exchange experiment, in which a powder sample exchanged at 0.17 h was dissolved in buffer of the same pH, gave no significant difference in exchange between powder and solution (Schinkel et al., 1985). (3) The 210-nm extremum of the circular dichroism spectrum of lysozyme changes by no more than 10% between dry and fully hydrated proteins (Chirgadze and Ovsepyan, 1972b). Because of contributions from surface chromophores, spectroscopic properties are difficult to interpret when comparing solution and powder conformation, but it is clear that drying does no violence to the chain folding. (4) The melting temperature increases strongly with decreasing hydration below 0.3 h (Fujita and Noda, 1978). Thus, there is no equilibrium or kinetic basis for expecting a conformation change with hydration. (5) Equivalence of conformation for

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the partially hydrated or dry protein and the fully hydrated protein is supported by other measurements, among which are the following: the IR spectrum changes smoothly with hydration above 0.07 h, and the changes are consistent with changes in the environment of surface hydrogen-bonding groups, without a change in conformation (Careri et al., 1979b);the partial specific volume of a protein is the solution value above about 0.15 h (Bull and Breese, 1968b; Richards, 1977); the order of amide hydrogen exchange is the same in lysozyme powders and solutions (Schinkel et al., 1985); onset of enzyme activity is 0.2 h for lysozyme and 0.12 h for chymotrypsin; some enzymes are active in nearly dry organic solvents (Klibanov, 1986; Zaks and Klibanov, 1984). Measurements such as the above collectively constitute support for the absence of hydration-dependent conformational rearrangements above the level of about 1 h;. Most measurements have been with small monomeric globular proteins. It is possible that larger or multimeric proteins are more sensitive to the removal of solvent. Are there effects of drying on the time-average conformation of a protein, other than changes within the 1 h; limit for backbone and buried side chains, discussed above? Apparently so. Drying changes the distribution of protons among the ionizable groups. Some spectroscopic properties show changes with hydration; these appear to be interpretable in terms of changes in surface side chains. The addition of solvent in molecular dynamics simulations leads to a general expansion of the molecule and some hydrogen bond rearrangement. X-Ray diffraction measurements for lysozyme suggest that there are domain displacements associated with drying. Morozov et al. (1988) analyzed the mechanical properties of protein crystals as a function of hydration; they proposed that the energy of mechanical deformation contributes importantly to the energetics of sorption and that there may be substantial associated changes in conformation of the individual protein molecules. Sections II-V described various data bearing on the comparison of protein in solution and in dry or partially hydrated samples. Most measurements have alternative interpretations, and it perhaps is good counsel to be wary of conclusions about conformation. VII. HYDRATION AND FUNCTION This section is a discussion of ways in which solvent can influence protein and enzyme processes. An understanding of these questions is just emerging, and little of what is presented here is as cleanly supported as the picture of hydration presented in the preceding section.

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A. Folding

Folding of a globular protein produces a surface that minimally affects the surrounding water. The thermodynamics of water in the interface differ only slightly from those of the bulk solvent, and the least possible amount of water is perturbed. This behavior can be seen as complementary to another aspect of protein folding: the withdrawal of hydrophobic side chains from solvent. The latter minimizes perturbation by burying those portions of the polypeptide for which water is the poorest solvent. The former minimizes perturbation of solvent by what remains exposed. Not all biological macromolecules show so small an effect. Nucleic acids require for their hydration about twice the amount of water required by globular proteins (for heat capacity measurements comparing protein and tRNA, see Rupley and Siemankowski, 1986). It may be significant that DNA, with an extensive hydration shell, underg,oesfacile hydration-dependent conformational transitions, which are not found for proteins. Domain coalescence (Karplus and Weaver, 1976) is a possible mechanism for protein folding. Zientara et al. (1980) examined the dependence of the coalescence lifetime on the hydration shell. The lifetime depends on the activation barrier contributed by the shell and the extent of the shell. If domains resemble the native protein in hydration, then the minimal extent of the shell and its fluidity favor coalescence. In passing, one notes that the percolation model may apply to folding: the coalescence of domains should be analogous to gelation or to diffusion on a partially filled lattice. Estimates of the contribution of hydration to the thermodynamics of folding have been based on additivity schemes and accessibility of elements of the protein to solvent (Eisenberg and McLachlan, 1986; Ooi and Oobatake, 1988a; Ooi et al., 1987).The methods merge macroscopic thermodynamic information about solutions of model compounds with an averaged molecular picture of the surface. Molecular dynamics (Levitt and Sharon, 1988) and Monte Carlo calculations (Hagler and Moult, 1978; Hermans and Vacatello, 1980)have given detailed pictures of protein-solvent interactions, allowing estimates of the thermodynamic differences between the hydration shell and the bulk solvent. Values obtained by the above methods agree qualitatively with experiments and expectation. Matthews and collaborators (see Alber and Matthews, 1987, and references cited therein) have used site-directed mutagenesis, thermodynamic measurements, and X-ray diffraction to develop an understanding of the folding thermodynamics and thermostability of phage T4

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lysozyme. Point mutations at an exposed residue (Thr-157) produced up to an 11" decrease in melting temperature. The analysis for one mutant (Thr- 157-Gly) suggested that a single water molecule, bound at position 157 and replacing the oxygen of Thr-157, stabilized the protein, in the sense that the mutation had a relatively small effect on melting temperature. The point is that individual water molecules at the protein surface may make significant contributions to the thermodynamics of folding. The relationship between the hydration shell and folding is of some importance for the use of enzymes as catalysts for syntheses, especially in industrial reactors (Wong, 1989). Nonaqueous media are preferred or are necessary for some reactions. Enzymes appear to be active when partially hydrated, at low water activity. In some cases there is activity in nearly dry nonaqueous solvents (Klibanov, 1986; Zaks and Klibanov, 1984). Thus, one should expect that, generally, it will be possible to find nonaqueous conditions for a particular enzyme-catalyzed process. Knowledge of the hydration shell is important, of course, for other aspects of the design of enzyme catalysts or drugs.

B . Chemistry of Transition States There is a considerable literature on the participation of solvent in reactions of small molecules. Bruice and collaborators systematically studied, as an enzyme model reaction, carboxylate attack on phenyl esters (see Bruice and Turner, 1970, and references cited therein). Rate enhancements of more than lo4were observed for some substrates upon transfer from water to dimethyl sulfoxide-water mixtures. The rates of nucleophilic displacement reactions can change by up to 20 orders of magnitude between gas phase and solution, and this effect has been related to the stepwise hydration of the nucleophile (Bohme and Mackay, 1981). Young and Jencks (1977)have shown that lysozyme must stabilize the putative oxocarbonium ion intermediate by 5-7 kcal/mol, compared with the species formed in the nonenzymatic hydrolysis of glycosides and acetals, which react rapidly with water. Bell et al. (1974) analyzed the reaction of chymotrypsin in several solvents, concluding that the substrate undergoes significant desolvation in the enzyme activation process. Warshel et d.(1989) pointed out that substrates are not desolvated on being bound by an enzyme; rather, the enzyme provides a special environment that does not resemble the gas phase desolvated state, but does serve to stabilize the transition state through the new interactions established.

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An expectation of work with powder samples is that analysis of the dependence of various properties on water activity will point to the role of hydration water. In the case of an enzyme, the most interesting would be its role in the rate-controlling step of the catalysis. The above remarks on chymotrypsin and lysozyme point out the fact that the environment of the transition state is likely to be dominated by protein groups. The hydration shell appears more likely to be involved in some other aspect of the catalysis: substrate binding; movement of reactants, including protons, about the enzyme surface; or coupling of protein motions. For example, the onset of activity for lysozyme is 0.2-0.25 h, which is closely correlated with two protein surface events. (1) Growth of the enzyme activity exactly parallels the hydration dependence of surface motion, detected with the ESR probe TEMPONE (Rupley et al., 1980). The lysozyme reaction is understood to involve movement of the substrate more deeply into the active-site cleft, during passage from the last equilibrium complex through the transition state of the rate-determining step to the oxocarbonium ion intermediate (Banerjee et al., 1975). Thus, development of enzyme activity in the partially hydrated powder, at the hydration level at which surface motion becomes unfrozen, is in accord with the mechanism of the catalysis, which requires mobility of the substrate on the enzyme surface. (2) The onset of protonic percolation for a lysozyme-substrate complex coincides with the onset of enzyme activity. Either long-range movement of protons may itself be important for a mechanism that comprises general acid catalysis or the surfacespanning hydrogen-bonded networks, which are first established at the percolation threshold, may have some other critical role. Percolation is discussed more fully in Section VI1,C. For other enzymes activity has been detected in some cases at 0.1 h or, rarely, at a lower hydration level. Most of the enzymes studied show onset of activity between 0.1 and 0.2 h, but there is not enough information to arrive at a consensus value. There appears to be no single hydration level that is critical for enzyme catalysis. Perhaps it is to be expected that different mechanisms should be associated with different roles of solvent. Despite the above generalizations, hydration water likely participates in the transition state for certain reactions. X-Ray diffraction results have identified, for some enzymes, bound solvent that may enter into the transition state, for example, lysozyme (Blake et al., 1983), DNase I (Oefner and Suck, 1986), and the aspartic proteinase from Rhizopus chinensis (Suguna et al., 1987). Meyer et al. (1988) suggested the participation of water organized in a network or tunnellike structure in the

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catalytic mechanism of serine proteases. Water threads and networks, generally seen in X-ray diffraction analyses of proteins, may be fundamental elements of enzyme catalytic structures, serving to couple protein processes. This possibility is discussed more fully below.

C . Protonic Conduction and Percolation Dielectric measurements (Careri et al., 1985, 1986) of partially hydrated powders of lysozyme have shown that protonic conduction is associated with a percolation transition involving water clusters. The threshold of the dielectric response is 0.15 h, corresponding to halfcoverage of the surface with water and to the percolation theory prediction, Binding of tetrasaccharide substrate at the active site reduces the proton flux by half. This would appear to mean that a significant proportion of the protonic conduction paths pass through the active site. The effect of substrate Binding is found at hydration levels near the end point, indicating that the special conduction paths persist at full hydration. Lysozyme, like many other enzymes, exhibits general acid-base catalysis as an element of its catalytic mechanism. It is an attractive possibility that the active site may be special in facilitating proton movement. The binding of tetrasaccharide shifts the hydration level of the percolation transition from 0.15 to 0.23 h. As noted in Section VII,B, the higher hydration level coincides with the onset of enzyme activity. Protonic percolation may be the event that imposes a lower limit on the hydration level for onset of enzyme activity, for those enzymes dependent on general catalysis. The hydration level at which chymotrypsin first displays activity, 0.12 h, is in agreement with this suggestion. The chymotrypsin mechanism includes general catalysis, but not significant substrate rearrangement in the rate-determining step. As noted, other enzymes show a critical hydration level between 0.1 and 0.2 h. For lysozyme both unfreezing of surface motion and protonic conduction correlate well with development of enzyme activity. It may be that both surface conduction and surface motion are essential to the operation of this enzyme. There are, however, other possibilities. Long-range proton movement, detected by the dielectric measurements, can be viewed as a probe of hydrogen-bonded networks. These appear at the percolation threshold and span all regions of the protein surface. Thus, any process tied to long-range connectivity would be established, as is protonic conduction, at the percolation threshold. Internal motions of the protein may be coupled to surface water networks (Doster et al., 1986). Surface motion also is plausibly tied to water arrangements suffi-

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ciently extensive to accommodate large groups-here, saccharide units. The percolation networks are stochastic, and they may be coupled with other statistical events of enzyme catalysis. The point is that enzyme catalysis, like other protein processes, is complex, and water may play several roles that are interdependent, possibly to such an extent that it may be inappropriate to try to separate them. D . Substrate Binding

Interactions between a substrate and the protein surface are perhaps 10 times fewer than the noncovalent interactions established in protein folding, yet the free energy of substrate binding is comparable to the free energy of protein folding. The special chemistry of the surface water may help to explain the tight binding of substrate, obtained through relatively few interactions with the protein, and the large free energies that have been associated with individual hydrogen bonds. The entropy of transfer of solvent into the interface is negative. Viewed most simply, if the entropy of the solvent at the surface reflects special chemistry, such as restrictions imposed by extended hydrogen-bonding arrangements, then displacement of this solvent should favor substrate binding. Also, crystallographic studies, such as those by Blake et al. (1983) on lysozyme, provide examples of water in the active site of an enzyme being located at the position occupied by a substrate atom in the enzyme-substrate complex. Again viewed most simply, this situation should favor substrate binding. Blevins and Tulinsky (1985) suggested two functions for the solvent at the chymotrypsin active site: (1) solvation of the Asp-His-Ser catalytic triad, and (2) a guiding effect on the substrate in formation of the enzyme-substrate complex, provided by several waters at the end of the specificity site. X-Ray diffraction results have suggested a role of activesite water in determining the kinetics or equilibria of substrate binding for other proteins (Section IV). Solution studies have addressed changes in hydration associated with substrate binding and catalysis. Haire and Hedlund (1983) found that high concentrations of ethylene glycol stabilized oxyhemoglobin. This was interpreted as an effect of change in protein hydration, seen as preferential binding of 5-10 mol of water per mol of hemoglobin. Sloan and Velick (1973) measured the buoyant density and preferential hydration of yeast glyceraldehyde-3-phosphate dehydrogenase and its complex with NAD. They concluded that there is a 15%decrease in preferential hydration, corresponding to a 6% volume contraction, when the

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coenzyme is bound. X-Ray diffraction studies have shown that conformational changes follow binding of cofactor and substrate to pyridine nucleotide-dependent dehydrogenases. Various authors have suggested, based on experimental observations, a deep involvement of change in hydration with movement along the enzyme reaction coordinate (Loftfield et al., 1980; Low and Somero, 1975a,b; Lumry and Gregory, 1986). Changes in the enzyme surface or its solvation, comparing the succession of ground states and transition states, can be understood to modulate the free energies of critical binding or activation steps at the expense of other segments of the reaction-in effect, the solvent functions as a means of free-energy transfer between reaction steps. Release of hydration water is an unfavorable process, and if it is to be an essential element of a mechanism, it should be through modulation of equilibria or activation events. A cooperative release of water could give a large cumulative effect, even though the free-energy change per water molecule transferred to bulk solvent may be small. An active site is typically several hundred square angstroms in area, covered by at least 10- 15 water molecules, perhaps more, if additional water in neighboring regions of the surface is included. E . Water Networks

Tapia and Eklund (1986) carried out a Monte Carlo simulation of the substrate channel of liver alcohol dehydrogenase, based on the X-ray diffraction structure for this enzyme. The addition of substrate and the associated conformation change induce an order-disorder transition for the solvent in the channel. A solvent network, connecting the active-site zinc ion and the protein surface, may provide the basis for a proton relay system. A molecular dynamics simulation of carbonic anhydrase showed two proton relay networks connecting the active-site zinc atom to the surrounding solvent (Vedani et al., 1989). They remain intact when the substrate, HCO, , is bound. Mackay and Wilson (1986) suggested that single-file waters in channel structures, like that studied in gramicidin A, may provide for long-range transfer of information, by a spatial (translational) response to a perturbation or by the correlated reorientation of water dipoles. Threads, clusters, or networks of water molecules have been detected by crystallographic, thermodynamic, and dynamic measurements. They appear to be a common feature of the protein-water interface, which should not be surprising, in view of the extensive hydrogen bonding of the bulk water. Moreover, the water of the interface differs from

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the bulk water, for example, in 10- 100 times slower motions and in the thermodynamics of transfer, and these differences may show up in the extended hydrogen-bonded arrangements of the surface. In any event hydrogen-bonded networks are linked, perhaps inseparably, with percolation processes, motions of the protein, and fluctuations. F. Fluctuations and Protein Motions

The concept of conformational substates serves to describe both timeaverage and dynamic aspects of protein conformation (for a review discussion see Frauenfelder et al., 1988). The minimum of the conformational energy surface corresponding to a thermodynamic state of a protein is understood to be rough, with valleys that define conformational substates and barriers that separate them. If the thermal energy is comparable to or greater than the barrier heights, a protein molecule will explore all of the substates within the envelope of the minimum in the energy surface. As the temperature is lowered, an individual protein molecule will be trapped in a particular substate, and an ensemble of molecules will be distributed among the traps, to give a glass. There may be a hierarchy of conformational substates, of mimina within minima, traversed successively as the temperature is lowered. This picture is supported by several lines of experimental evidence, two principal ones being Mossbauer spectroscopic and other measurements of the transition near 200 K (Sections 111, IV, and VI), believed to correspond to formation of a glassy state, and low-temperature X-ray diffraction measurements (Section IV), which show appreciable disorder remains after large-scale collective motions have been frozen out. The hydration water plays two roles in this picture. (1) It serves to catalyze transitions between substates that involve changes in hydrogen bonding, through participating in alternative hydrogen bond arrangements that conserve the total number of hydrogen bonds in the system (Chirgadze and Ovsepyan, 1972a). (2) It serves to couple fluctuations of the protein-water system. This is discussed in the previous section, which lists evidence for solvation water-dependent coupling and the elementary mechanisms that might operate. G . Fluctuations and Catalysis We start from the experimental observation that lysozyme powders exhibit a single critical hydration level for the onset of enzyme activity and the onset of surface motions, displayed in the dynamics of a spin

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probe and in protonic conduction. The critical hydration level can be viewed as the necessary condition for spontaneous fluctuations in the protein-water structure, which, in turn, can be viewed as essential for function. This hydration level is also the point at which long-range connectivity among the surface water molecules is established, allowing proton displacement along statistical paths over the surface of the enzymesubstrate complex. An attractive inference is that the role of hydration in lysozyme function is to provide both for structural adjustmentsnamely, the sequence of isomerizations that define the sequential structure of the reaction pathway-and for the long-range charge displacements that may be coupled to passage through the transition state. This statistical picture of the role of protein hydration may also apply to other proteins. From a statistical viewpoint enzyme action appears as a series of programmed events that take place in a thermal bath. Careri (1974) proposed the relevance for enzyme action of cross-correlated fluctuations in the Onsager formulation. Suggestions have been made concerning the coupling of protein fluctuation events and catalysis (Careri and Gratton, 1986). The hydration shell must play a role, if only in coupling enzyme fluctuations with the surrounding thermal bath (Careri and Gratton, 1977). A succession of rearrangements (i.e., isomerizations) of the geometry of the hydrated enzyme-substrate complex is perhaps a universal aspect of enzyme reactions. The hydration shell should facilitate optimization of the active-site environment during traversal of the reaction path. Again, mechanisms of coupling mediated by the hydration shell were discussed in the previous section. The conformational substate picture of protein dynamics just described has been applied to protein function. Two classes of motion were distinguished: equilibrium fluctuations, which are between substates within the envelope of a single thermodynamic state of a protein, and functionally important motions, which are not governed by equilibrium thermodynamics and may involve larger changes in the protein conformation. The view of the coupling between the two classes of motion is essentially that given in the preceding paragraphs. For a discussion see Frauenfelder and Gratton (1986).

H . Membranes There should be similarities between proteins at low hydration and proteins that are in membranes and consequently have a portion of their surface withdrawn from solvent. Some water may be associated with the

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protein surface internal to the membrane (Rupley and Siemankowski, 1986). Dielectric measurements on the purple membrane of Halobucterium halobium (Rupley et al., 1988), similar to those carried out for lysozyme, have demonstrated two-dimensional protonic percolation, with somewhat more complex properties than those found for lysozyme. The percolation threshold is at the hydration level at which changes in the photoresponse have been observed (Varo and Keszthelyi, 1983). The percolation model suggests that it may not be necessary to have a rigid geometry and definite pathway for conduction, as implied by the “proton-wire” model of membrane transport (Nagle and Mille, 1981). For proton pumps the fluctuating random percolation networks would serve for diffusion of the ion across the water-poor protein surface, to where the active site would apply a vectorial “kick.” In this view the special nonrandom structure of the active site would be limited in size to a dimension commensurate with that found for active sites of proteins such as enzymes. Control is possible: conduction could be switched on or off by the addition or subtraction of a few elements, shifting the fractional occupancy up or down across the percolation threshold. Statistical assemblies of conducting elements need only partially fill a surface or volume to obtain conduction. For a surface the percolation threshold is at half-saturation of the sites. For a three-dimensional pore only onesixth of the sites need be filled. Hydration forces, the long-range interactions between surfaces in water, may participate in membrane function, specifically, in the response to diacylglycerol, in membrane fusion, and in the gating of membrane channels (Rand et al., 1985; Zimmerberg and Parsegian, 1987).

I. Complex System The effect of solvent on rates and thermodynamics of reactions can be understood to propagate through macromolecular processes to an influence of solvent on higher levels of organization. Biological systems are, by definition, multicomponent systems. One should keep in mind the difficulties of constructing molecular level pictures that satisfactorily describe systems such as a protein in a reverse micelle or a protein in a concentrated aqueous salt solution, which are certainly much simpler than anhydrobiotic organisms, for example. It is not clear to what extent the water of the hydration shell can be replaced by a third component (e.g., lipid) or what effect such replacement has on protein or enzyme properties.

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I . Anhydrobiosis A variety of biological systems, termed “anhydrobiotic” or “cryptobiotic,” exist at low water activity. These include seeds, spores, and cysts or desiccated forms of organisms such as the brine shrimp, Artemia, and certain nematodes. For review discussions, including summaries of physical and biochemical measurements made on these systems, see Leopold (1986) and Crowe and Clegg (1973, 1978). There are substantial changes in metabolism associated with generation of the desiccated state. Some organisms produce saccharides that serve to maintain the integrity of membranes. Also, desiccation is a trigger to switch the metabolism of seeds from a developmental to a germination mode. The water content of an anhydrobiotic organism is a function of the relative humidity of the environment. Under typical conditions it is 0.1 h for seeds or Artemza cysts. At this hydration level there is essentially no metabolic activity. 2. Antifreeze Proteins

Polar or subpolar fish survive at - 1.9”C,the freezing temperature of sea water. The blood of these fish contains compounds that depress the equilibrium freezing point by about 1°C. The blood also contains proteins that further lower the freezing point by reducing the rate of growth of ice nuclei (DeVries, 1984, 1986; Feeney et al., 1986). These compounds are of two main classes: antifreeze glycoproteins (AFGPs), with a repeating Ala-Ala-Thr(disaccharide) sequence, and antifreeze proteins (AFPs), which are rich in alanine but also contain a variety of other amino acids. Ice nuclei grow most rapidly in a direction orthogonal to the c axis of ice I. Both AFGPs and AFPs reduce the freezing point by binding to the growth plane and blocking growth (Raymond and DeVries, 1977). The AFGP interaction appears to be through a specific fit of the saccharide hydroxyl groups to the ice surface. An AFGP has been shown to inhibit the melting of single crystals of pure ice, an activity symmetrical to the blocking of crystal growth (Knight and DeVries, 1989).Recently, the crystal structure of an AFP was determined at 2.5 h; resolution (Yang et al., 1988). The 37-residue polypeptide is a single a helix. There appears to be no plausible and unique matching of distances between side chains of the protein and the ice lattice. Yang et al. (1988) suggested that interaction of the helix dipoles with the ice accounts for the alignment of the AFP, with side-chain interactions with the ice serving to stabilize the complex. In any event the AFPs present a clear example of the functional importance of the protein-solvent (here,

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ice) interface, which may be relevant for understanding the proteinwater interface. Proteins from certain bacteria are ice nucleators (Wolber et al., 1986). An AFGP inhibits this nucleation, apparently by binding to the bacterial protein. 3 . Food

The properties of foods areinfluenced by water content and hydration level in several ways: texture, as for doughs; storage life, as affected by enzymatic and nonenzymatic reactions and by microbial growth; ease and cost of processing, as in the energy required for freezing or dehydrating a food product; effects of high or freezing temperatures; and marketing and appearance (for review discussions see Duckworth, 1975, and Simatos and Multon, 1985). The hydration dependence of food properties is linked to the hydration of proteins, both through proteins’ being a major constituent of foods and, more important, through the enzyme reactions governing food maturation and degradation. VIII. CONCLUSION A proper response to the question (Maddox, 1988) “Is the scandal, that so little is known about the interactions of macromolecules and their aqueous environment, about to be removed?” appears to be “Yes, for proteins.” The description of the interface between protein and solvent is reasonably satisfactory. On a scale of 1-10, it might receive a grade of 8. Perhaps the principal motive for devoting attention to the interface is the expectation that it contributes importantly to protein function, particularly enzyme function. From the preceding section it is clear that the correspondences that can be drawn now are, more often than one would like, speculative, worth no more than a grade of perhaps 4 for identification of the ways in which the hydration shell might participate in function, and at best a grade of 2 for the quantitative understanding of ways in which it does participate. However, the directions in which to move for a more complete analysis of the coupling of hydration and function are becoming clear. In cases in which a sharp question can be formulated and so suggest a test of a mechanism of solvent participation, site-directed mutagenesis provides an experimental tool, perhaps appropriately guided by computer simulation of the protein-solvent system. The participation of fixed or random (percolative)network structures in proton movement is

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a particularly attractive possibility for a function of surface water, and suggestions of this kind appear to be open to testing. Experimental observations, such as the coincidence of onset of function with onset of protein motion and long-range connectivity, lead one to infer that correlated fluctuations are among the fundamental principles of enzyme activity. The proof of this inference, through the detection of cross-correlations,remains an open and difficult problem. There is a substantial body of evidence, however, that bears on the correlation of solvent and protein motions, and understanding of the details of this coupling is likely to come soon. Connectivity has a central role in biochemistry and biology, and one imagines that the percolation model, with its focus on connectivity, should have wide application. Percolative behavior is to be expected for the coordinate functioning of systems of proteins in metabolic pathways, for functional interactions between proteins embedded in a membrane, for the interactions between domains in the folding of a polypeptide, or for the onset of function in anhydrobiotic organisms, seeds, and spores. Solvation of proteins in multicomponent systems is associated with interesting chemistry and has practical applications. Clever experiments have been carried out in this area, but there as yet has been no systematic exploration of the chemistry, such as is needed for development of a working picture of a protein in a multicomponent environment, particularly the complex environments of membranes, organelles, and cells. There are several different, but not mutually exclusive,pictures of protein hydration. The surface picture, emphasized in this review, is appropriate for much previous and current work on hydration. It is based on the interaction of water molecules with protein surface sites, it deals only with relatively strong perturbations of the solvent and protein, and it is simple to model. The picture for electrostatic interactions treats the solvent as a continuum. The picture for hydration forces treats long-range and weaker protein-water interactions and emphasizes solvent-solvent interactions and cooperative effects. Other pictures deal with protein hydrodynamics and preferential binding. One expects that several of these seemingly different views can be merged, perhaps without great difficulty, into a comprehensive model of hydration. In this regard computer simulations are being applied with greater frequency to the description of protein solvation. Their contributions to the refinement of molecular-level pictures should grow to be even more significant as potential functions are improved and machines become faster. Many of the current problems appear to be accessible via experimen-

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tation and theory, so ensuring the continuing vigorous evolution of the understanding of hydration. APPENDIX: PERCOLATION THEORY A. Introduction

The term “percolation” is widely used and is reminiscent of the notion of diffusion in a stochastic medium that is only partially connected, such as a porous material. Broadbent and Hammersley introduced percolation theory in 1957, to describe vapor flow in a partly blocked filter. The original use of the methods of percolation theory is due, however, to Flory (1941) and Stockmayer (1943), whose work was recognized only much later as an application of percolation theory. During World War I1 they described how small branching molecules form increasingly larger macromolecules as more chemical bonds are formed. This polymerization process may lead to gelation, as in the boiling of an egg, and biochemists may be experienced in percolation without being aware of it. The power of percolation theory is its offering of a unitary treatment for widely different phenomena occurring in disordered systems. The central feature of percolation theory is the existence of a threshold at which long-range connectivity suddenly emerges. The following simple experiment demonstrates the essential concept of the critical threshold for percolation. A mixture of small plastic and metal balls of equal size is poured into a beaker with a crumpled-foil electrode at the bottom, another crumpled-foil electrode is pressed onto the top, and the electrodes are connected to a battery through an ammeter. Current is measured as a function of the composition (i.e., fraction of metal balls) of the conductorlinsulator mixture. There is a critical composition below which no current flows and above which the conductivity increases nearly exponentially. At this threshold the two electrodes suddenly become spatially connected along a statistical pathway originating in the random medium. Percolation theory tells us that the critical composition is 0.25 fraction metal balls, a remarkably low concentration. This is perhaps not an intuitive result. A second tutorial example may be useful. Suppose that an extended communication network, modeled as a large two-dimensional squarelattice grid connected to heavy bars at two opposite boundaries, is attacked by a stochastic saboteur, who, with wire-cutters, severs the grid interconnections. What fraction of the links must be cut in order to isolate the two bars from each other? The answer, given by a a percolation

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analysis, is 0.5. There are two important points: first, the critical composition depends on the dimensionality of the system; second, the sharp critical transition has as its essential feature the switch between the presence and absence of long-range connectivity, specifically, the “infinite” or “lattice-spanning”or “unbounded” cluster, which, above the percolation threshold, embraces all regions of the space. The percolation model can be stated more precisely than was done above. A lattice is composed of sites (vertices, or intersections between bonds) and bonds (links, or connections between sites). There are two basic types of percolation processes on lattices: bond percolation and site percolation. In bond percolation each bond is either connected (with probability p ) or disconnected [with probability (1 - p)]. In site percolation each bond is considered to be connected, and it is now the sites which carry the random connectivity of the structure, each site being filled (with probability p ) or empty [with probability (1 - p ) ] . A set of connected bonds or sites is called a cluster. In the following we shall assume the probability p to be the same for each site (or bond) and not to be influenced by the state of the neighboring siteslbonds. This is equivalent to assuming that there is no energy of interaction between connected sitedbonds. There are more complex percolation models. For example, in site-bond percolation the composition of the system is specified by two independent variables, p(site) and p(bond). In order for there to be a percolation path, both p(site) and p(band) must be large, and how large each must be depends on the other; namely, they are interdependent. Site and bond percolation can be shown to be equivalent, and for simplicity from this point we discuss only site percolation. The percolation threshold p , is defined as the site-filling probability that marks the appearance of the lattice-spanning percolation cluster and the establishment of long-range connectivity. One can introduce the function P ( p ) , called the percolation probability, which has the following significance: When the fraction of filled sites is p , P ( p ) is the chance of a randomly chosen site being both filled and part of the infinite cluster, or, in other words, P ( p ) is the fraction of the entire system that is taken up by the infinite cluster. P is the key function characterizing a percolation process, and here it plays the role of the order parameter used to describe order-disorder phenomena and phase transitions. Its behavior for a square lattice is show in Fig. 39. Percolation theory provides a well-defined model applicable to a wide variety of spatially random phenomena, both macroscopic and microscopic (Table X). The characteristic length scales for these phenomena

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TABLE X Phenomena Modeled

h Percolation Them

Phenomenon or system

Transition

Flow of liquid in porous medium Spread of disease in population Communication or resistor networks Conductor-insulator composite materials Composite superconductor-metal materials Discontinuous metal films Stochastic star formation in spiral galaxies Quarks in nuclear matter Thin helium films on surfaces Metal atom dispersions in insulators Dilute magnets Polymer gelation, vulcanization Glass transition Mobility edge in amorphous semiconductors Variable-range hopping in amorphous semiconductors

LocaUextended wetting Containment/epidemic Disconnected/connected Insulator/metal Normalhperconducting Insulatorhnetal Nonpropagation/propagation Confinement/nonconfinernent Normal/superfluid Insulator/metal Paramagnetic/ferromagnetic Sol/gel Liquid/glass Localized/extended states Resistor network analog

From Deutscher et al. (1983).

differ by many orders of magnitude, but the properties in the vicinity of the percolation threshold are similar for all of them. The interested reader may want to consider several fine reviews (e.g., De Gennes, 1976; Essam, 1980), the collection of articles in the volume Percolation Structures and Processes (Deutscher et al., 1983), and the excellent books by Zallen (1983) and by Stauffer (1985). In the following parts of this section, we consider only those concepts used in this review. €3. Invariant Quantities

Although percolation theory deals with random systems, modeling and numerical calculations for percolation are usually carried out for a lattice of some definite geometry. To reach conclusions which do not depend on the details of lattice geometry but on dimensionality only, and thus are valid for the random system of interest, some invariant quantities must be constructed. One such invariant is the critical volume fraction for percolation, introduced by Scher and Zallen (1970). The critical volume fraction +(d) may be defined as the average (p,,(d) x v,(d)), where p,,(d) is the percolation threshold for simple (nearest-neighbor sites only) lattice j in d dimensions, and vj(d) is the filling factor of the lattice, corresponding to the packing of equal, touching, and nonoverlapping d-dimensional spheres, centered on lattice sites. The utility of 4(d)is based on the em-

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pirical rule that in both two and three dimensions &d) is nearly independent of the lattice type j for given d; namely, it is a dimensional invariant. This statement is explained better by examination of Table XI (compiled from the data of Zallen, 1983). Thus, for the random close-packed structure of conducting and nonconducting spheres, considered above, for which the filling factor II = 0.637, the percolation threshold p, is simply calculated from the lattice-invariant three-dimensional percolation threshold value, $(d = 3) = 0.16, as being

pc = 0.1610.637 = 0.25 It is clear that the introduction of a critical volume fraction is a step toward dealing with percolation on a continuum. To this end Zallen and Scher (1971) considered the motion of a classical particle in a random potential, V ( r ) ,and introduced a function, + ( E ) , which defines the fraction of space accessible to particles of energy E. The connection with percolation is in the fact that, for energies such that &E) > &(E), there are infinitely extended volumes of allowed ( V < E) space. The critical value $c is identified with 0.15 for d = 3, and “delocalized states” appear above + c . Let us move now to the important region close to the percolation threshold, namely, the critical region where ( p - p,l << 1. In this region the percolation functions are observed to obey power law dependences on the distance from the threshold lp - pcI. Thus, for the divergence of the average size S,, and linear dimension 1,” of the clusters, as p, is approached from below, we write =

14pc - p ) . W p c - p).

& I ”

saw

=

where Y and y are the critical exponents, as given in Table XII. Now the remarkable fact is that, in spite of the great dependence of p , on the lattice type (see Table XI), these critical exponents do not depend on the details of lattice geometry and are the same for all lattices of the same dimensionality. Values of critical exponents for the percolation functions already introduced and for the conductivity are given in Table XI1 (data from Stauffer, 1985). Note that no distinction has been made between site percolation and bond percolation in Table XII. This is a further tribute to the generality of the critical exponents; in the jargon of phase transitions, site and bond percolation are said to belong to the same “universality class.” Admittedly, percolation displays an unusual phase transition, in that it is temperature insensitive. It is a true phase transition, however, with many functions diverging or vanishing at one sharply defined point, the percolation threshold pc. In percolation theory these

TABLE XI Critical Concentrations for Bond Dimensionality

(4

Lattice or structure

(pC6Ond) and Site ( p f ” ) Percolation on Lattices a Coordination

pcbond

PPte

Filling factor (v)

(z)

zx

pcbond

4c = u x p2‘r

~

1 2

Chain Triangular Square Kagomk Honeycomb

1 0.3473 0.5000 0.4500 0.6527

1 0.5000 0.5930 0.6527 0.6980

I 0.9069 0.7854 0.6802 0.6046

1 0.45 0.47 0.44 0.42

2 2.08 2.00 1.80 1.96 ~

2.0 3

fcc bcc sc Diamond

0.119 0.179 0.247 0.388

0.198 0.245 0.311 0.428

12 8 6 4

0.7405 0.6802 0.5236 0.3401

“fcc, Face-centered cubic; bcc, body-centered cubic; sc, synclinal. (From Deutscher et al., 1983.)

5

0.2

0.45

2

1.43 1.43 1.48 1.55

0.147 0.167 0.163 0.146

1.5 f 0.1

0.16

If-

0.03

0.02

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TABLE XI1 Percolation Exponents for Two and Three Dimensions, and Corresponding Percolation Quantity a

Exponent

d =2

d =3

Quantity

P

5/36 43/18 413 1.3

0.40 1.80 0.90 2.00

Percolation probability Mean size of finite clusters Correlation length Conductivity

Y V

U

"d, Dimension. (From Deutscher et al., 1983.)

functions are purely geometric quantities, but the similarity between the more familiar types of phase transition and the percolation phase transition becomes clear when we consider the scaling laws governing asymptotic behavior close to the critical point. The values of the exponents quoted in Table XI1 have been estimated numerically by renormalization group techniques. Intuitively, there should be a close relationship between conductivity and percolation probability, and one would guess that their critical exponents should be identical. This is not true. Dead ends contribute to the mass of the infinite network described by the percolation probability, but not to the electric current it carries. Figure 39 shows the different growth of the percolation probability and the conductivity. It is convenient to set the conductivity equal to unity at p = 1, as in Fig. 39. We note, in passing, that diffusivity is proportional to conductivity, in agreement with Einstein's result in statistical physics: that diffusivity is proportional to mobility. 1

Oo PC

FIG. 39. Behavior of the percolation probability, P, and the conductivity, u,as a function of the concentration of filled sites, p . From Zallen (1979).

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JOHN A. RUPLEY AND GIORGIO CARER1

C . Finite-Size Effects The numerical results reviewed above were obtained for infinite lattices. How do the various quantities of interest behave near the percolation threshold in a large but finite lattice?This problem has been studied by renormalization methods, which are essentially equivalent to finitesize scaling. For finite lattices the percolation transition is smeared out over a range of p , and one must expect a similar trend in other functions, including the conductivity. Computer simulations by the Monte Carlo method have been carried out for bond percolation on a threedimensional simple cubic lattice by Kirkpatrick (1979). Five such “experimental” curves are shown in Fig. 40, each of which corresponds to a cube of size b, containing bS bonds. In Fig. 40 the vertical axis gives the fraction p‘ of such samples that percolate (i.e., have opposite faces conI.0-

0.0

-

0.6

.-

p’

IFRPtTlON OF ClMUCTlNG SPMPLES 1 0.4

-

-

0.2

0

0.20

0.22

p

0.24

0.26 0.28 MKSENT 1

0.30

( FRACTION OF BONOS

FIG.40. Scaling for finite-sized lattices. Computer calculations of scaling properties for bond percolation on the three-dimensional simple cubic lattice. When p is the fraction of connected bonds, p’ = p ’ ( p ,b) is the fraction of cubic samples of edge length b that contain a continuous path of connected bonds (a spanning cluster) which links opposite faces of the sample. From Kirkpatrick (1979).

161

PROTEIN HYDRATION AND FUNCTION

1.u

.8.

I

4P) .4’

I

r

ff P

t

0

0

.2

.6

.4

.0

1.0

P-

FIG. 4I. Percolation probability for finite-sized lattices. Computer calculations of the percolation probability, P ( p ) , as a function of the site-filling probability, p, for twodimensional square lattices of varied dimension: 0, 10 X 10; 0 , 20 X 20; *, 40 X 40. Each curve is an average over a set of site percolation simulations for a lattice size. The site percolation threshold for an infinite two-dimensional square lattice is 0.593. Nonzero values of P ( p ) below the infinite lattice threshold reflect the variance of the threshold value for finite lattices (unpublished results).

nected). In the limit of an infinite lattice, the function p ’ ( p ) becomes a step function that jumps from p’ = 0 to p’ = 1 at the percolation threshold p , . Similar finite-size effects are found for two-dimensional samples. Figure 41 shows the percolation probability P ( p ) , determined by averaging Monte Carlo simulations for site percolation on a twodimensional square lattice, for finite lattices of varied size. For an infinite lattice of this type, p, = 0.593. The nonzero values of P ( p ) below p = 0.593 reflect the dispersion in p , found for finite lattices. A protein, with several hundred water sites on its surface, would fall in the range of lattice sizes modeled in Fig. 41. The shape of the P ( p ) function is not strongly affected by lattice size.

REFERENCES Abraham, F. F. (1974). “Advances in Theoretical Chemistry, Suppl. 1: Homogeneous Nucleation Theory. The Pretransition Theory of Vapor Condensation.” Academic Press, New York.

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Acker, L. (1962). Adv. Food Res. 11,263-330. Adair, G. S., and Adair, M. E. (1936). Proc. R. Sac. London, Ser. B 120,422-446. Ahlstroem, P., Teleman, O., Joensson, B., and Forsen, S. (1987). J. Am. Chem. Sac. 109, 1541 - 155 1. Alber, T., and Matthews, B. W. (1987). In ‘:Protein Engineering” (D. L. Oxender and C. F. Fox, eds.), pp. 289-297. Liss, New York. Ali, S., and Bettelheim, F. A. (1985). Colloid Polym. Sci. 263,396-398. Aliotta, F., Fontana, M. P., Giordano, R., Migliardo, P., and Wanderlingh, F. (1981). J . Chem. Phys. 75,4307-4309. Almog, R., and Schrier, E. E. (1978).J. Phys. Chem. 82,1701-1702. Anagnostopoulou-Konsta, A., and Pissis, P. (1987).J . Phys. D: Appl. Phys. 20, 1168- 1174. Andrew, E. R. (1985). Polymer 26, 190- 192. Andrew, E. R., Bryant, D. J., and Rizvi, T. Z. (1983). Chem. Phys. Lett. 95,463-466. Andronikashvili, E. L., Mrevlishvili, G. M., Yaparidze, G. S., and Sokhadze, V. M. (1979). Int. J . Quantum Chem. 16, 367-377. Ataka, M., and Tanaka, S. (1979). Biopolymers 18,507-516. Ataka, M., and Tanaka, S. (1980). Biopolymers 19,669-679. Aviram, I., and Schejter, A. (1972). BiopOly~m511, 2141-2145. Baker, E. N., and Hubbard, R. E. (1984). Prog. Biophys. Mol. Biol. 44,97-179. Baker, E. N., Blundell, T. L., Cutfield, J. F., Cutfield, S. M., Dodson, E. J., Dodson, G. G., Hodgkin, D. M. C., Hubbard, R. E., Isaacs, N. W., Reynolds, C. D., Sakabe, K., Sakabe, N., and Vijayan, N. M. (1988). Philos. Trans.R . Sac. London, Ser. B 319, 369456. Baker, L. J., Hansen, A. M. F., Rao, P. B., and Bryan, W. P. (1983). Biopolymers 22, 1637-1640. Baker, T., Dodson, E., Dodson, G., Hodgkin, D., and Hubbard, R. (1987). NATO Adv. Study Znst. Ser., Ser. A 126, 179- 192. Banerjee, S. K., Holler, E., Hess, G. P., and Rupley, J. A. (1975). J. Biol. Chem. 250, 4355-4367. Barbaric, S., and Luisi, P. L. (1981).J. Am. C h m . Sac. 103,4239-4244. Barlow, D. J., and Poole, P. L. (1987). FEBS Lett. 213,423-427. Barlow, D. J., and Thornton, J. M. (1988).J. M o l . Biol. 201,601-619. Bash, P. A., Singh, U. C., Brown, F. K., Langridge, R., and Kollman, P. A. (1987). Science 235,574-576. Bauminger, E. R., Nowik, I., Harrison, P. M., and Treffry, A. (1987). Hyperfine Interact. 33, 241 -248. Beece, D., Eisenstein, L., Frauenfelder, H., Good, D., Marden, M. C., Reinisch, L., Reynolds, A. H., Sorensen, L. B., and Yue, K. T. (1980). Biochemistry 19,5147-5157. Behi, J., Bone, S., Morgan, H., and Pethig, R. (1982). Int.J. Quantum Chem., Quantum Biol. Symf. 9,367-374. Bell, R. P., Critchlow, J. E., and Page, M. I. (1974).J . C h . Sac., Perkin Trans.2, 66-70. Belonogova, 0. V., Frolov, E. N., Krasnopol’skaya, S. A., Atanasov, B. P., Gins, V. K., Mukhin, E. N., Levina, A. A., Andreeva, A. P., Likhtenshtein, G. I., and Gol’danskii, V. I. (1978). Dokl. Biophys. (Engl. Traml.) 241,219-222. Belonogova, 0. V., Frolov, E. N., Illyustrov, N. V., and Likhtenshtein, G. I. (1979). Mol. Biol. (Moscow)13,567-576. Ben-Naim, A., Ting, K. L., and Jernigan, R. L. (1989a). Biopolymers 28, 1309-1325. Ben-Naim, A., Ting, K. L., and Jernigan, R. L. (1989b). B i o p o l p r s 28, 1327-1337. Berlin, E., K h a n , P. G., and Pallansch, M. J. (1970). J. Colloid I n t 4 a c e Sci. 34,488-494. Bernhardt, J., and Pauly, H. (1975).J. Phys. Chem. 79,584-590. Bernhardt, J., and Pauly, H. (1980).J. Phys. Chem. 84, 158-162. Betzel, C., Pal, G. P., and Saenger, W. (1988). Eur.J. Biochem. 178, 155-171.

PROTEIN HYDRATION AND FUNCTION

163

Blake, C. C. F., Pulford, W. C. A., and Artymiuk, P. J. (1983). J . Mol. Biol. 167, 693-723. Blevins, R. A,, and Tulinsky, A. (1985).J. Biol. Chem. 260,8865-8872. Bohme, D. K., and Mackay, G. I. (1981).J. Am. Chem. SOC.103,978-979. Bone, S. (1987). Biochim. Biophys. Acta 916, 128-134. Bone, S., and Pethig, R.‘(1982).J.Mol. Biol. 157,571-575. Bone, S., and Pethig, - R. (1985). 1. Mol. Biol. 181,323-326. Bone, S., Eden, J,, and Pethig, R. (1981). Int. J. Quantum Chem., Quantum Biol. Symp. 8, 307-3 16. Borah, B., and Bryant, R. G. (1982). Biophys. J . 38,47-52. Bourret, D., and Parello, J. (1984).J . Phys., Colloq. (Orsay, Fr.), 255-258. Bridelli, M. G., Capelletti, R., Ruani, G., and Vecli, A. (1985). Proc.-Int. Symp. Electretc, 5th, 831-835. Brooks, B., and Karplus, M. (1985). Proc. Natl. Acad. Sci. U.S.A. 82,4995-4999. Brooks, C. L., 111, and Karplus, M. (1986). In “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 369-400. Academic Press, Orlando, Florida. Brooks, C. L., 111, and Karplus, M. (1989).J. Mol. Biol. 208, 159-181. Brown, K. A., Brick, P., and Blow, D. M. (1987). Nature (London) 326,416-418. Brown, R. D., 111, and Koenig, S. H. (1982). Biophys. Chem. 16, 133-137. Bruccoleri, R. E., Karplus, M., and McCammon, J. A. (1986). Biopolymers 25, 1767-1802. Bruice, T. C., and Turner, A. (1970).J . Am. Chem. SOC.92,3422-3428. Brunauer, S., Emmett, P. H., and Teller, E. (1938).J. Am. Chem. SOC.60,309. Bryan, W. P. (1980).J. Theor. Biol. 87,639-661. Bryan, W. P. (1987). Biopolymers 26, 1705-1716. Bryant, R. G. (1978). Annu. Rev. Phys. Chem. 29, 167-188. Bryant, R. G. (1988). Stud. Phys. Theor. Chem. 38,683-705. Bryant, R. G., and Shirley, W. M. (1980a). Biophys.J. 32,3-16. Bryant, R. G., and Shirley, W. M. (1980b). ACS Symp. Ser. 127, 147-156. Bryant, R. G., Brown, W. E., 111, Sutcliffe, J. W., and Pulsinelli, P. D. (1983). Biochemistr)l 22,2914-2923. Bull, H. B., and Breese, K. (1968a). Arch. Biochem. Biophys. 128,488-496. Bull, H. B., and Breese, K. (1968b). Arch. Biochem. Biophys. 128,497-502. Careri, G. (1974). Stud. Nat. Sci. (N.Y.) 4, 15-35. Careri, G., and Gratton, E. (1977). BioSystems 8, 185-186. Careri, G., and Gratton, E. (1986). In “The Fluctuating Enzyme” (G. R. Welch, ed.), pp. 227-262. Wiley, New York. Careri, G., Angelis, L. D., Gratton, E., and Messana, C. (1977). Phys. Lett. A 60A,490-491. Careri, G., Fasella, P., and Gratton, E. (1979a). Annu. Rev. Biophys. Bioeng. 8, 69-97. Careri, G., Giansanti, A., and Gratton, E. (1979b). Biopolymers 18, 1187-1203. Careri, G., Gratton, E., Yang, P. H., and Rupley, J. A. (1980). Nature (London) 284, 572-573. Careri, G., Geraci, M., Giansanti, A., and Rupley, J. A. (1985). Proc. Natl. Acad. Sci. U.S.A. 82,5342-5346. Careri, G., Giansanti, A., and Rupley, J. A. (1986). Proc. Natl. Acad. Sci. U.S.A. 83, 6810-6814. Careri, G., Giansanti, A., and Rupley, J. A. (1988). Phys. Rev. A 37,2703-2705. Caspar, D. L. D., Clarage, J., Salunke, D. M., and Clarage, M. (1988). Nature (London) 332, 659-662. Cavatorta, F., Fontana, M. P., and Vecli, A. (1976).J. Chem. Phys. 65, 3635-3640. Celaschi, S., and Mascarenhas, S. (1977). Biophys. J. 20,273-277. Cerofolini, G. F., and Cerofolini, M. (1980).J. Collozd Interface Sci. 78,65-73. Chirgadze, Y. N. (1972). Dokl. Biophys. (Engl. Traml.) 204, 723-726. Chirgadze, Y. N., and Ovsepyan, A. M. (1972a). Biopolymers 11,2179-2186,

164

JOHN A. RUPLEY AND GIORGIO CARER1

Chirgadze, Y. N., and Ovsepyan, A. M. (1972b). Mol. B i d . (Moscow) 6,721-726. Churg, A. K., and Warshel, A. (1986).Bkchemistty 25,1675-1681. Claesson, P. M., Arnebrant, T., Bergenstaahl, B., and Nylander, T. (1989).J . Colloid Interface Sci. 130,457-466. Clementi, E. (1985).J . P h y . Chem. 89,4426-4436. Clementi, E., and Chin, S., eds. (1986). “Structure and Dynamics of Nucleic Acids, Proteins, and Membranes.” Plenum, New York. Clementi, E., Ranghino, G., and Scordamaglia, R. (1977). Chem. Phys. Lett. 49, 218-224. Clerc, J. P., Giraud, G., Alexander, S., and Guyon, E. (1980). Phys. Rev. B : Condens. Matter 22,2489-2494. Crowe, J. H., and Clegg, J. S., eds. (1973). “Anhydrobiosis.”Dowden, Hutchinson and Ross, Stroudsburg, Pennsylvania. Crowe, J. H., and Clegg, J. S., eds. (1978). “Dry Biological Systems.” Academic Press, New York. Cusack, S. (1986). Comments Mol. Cell. Btqphys. 3,243-271. Cusack, S. (1989).NATO Adv. Study Inst. Ser., Ser. A 178, 103-122. Cusack, S., Smith, J., Finney, J., Karplus, M., and Trewhella, J. (1986). Physzca B + C (Amsterdam) 136,256-259. Cusack, S . , Smith, J., Finney, J., Tidor, B., and Karplus, M. (1988). J . Mol. Biol. 202, 903-908. De Gennes, P. G. (1976). Recherche 7,919. Delepierre, M., Dobson, C. M., Howarth, M. A., and Poulsen, F. M. (1984).Eur.J. Biochem. 145,389-395. Deutscher, G., Zallen, R.,and Adler, J., eds. (1983). “Annals of the Israel Physical Society, Vol. 5: Percolation Structures and Processes.” Hilger, Bristol, England. Deutschmann, G., and Ullrich, V. (1979).Anal. Biochem. 94,6- 14. DeVries, A. L. (1984). Philos. Trans. R. Soc. London, Ser. B 304,575-588. DeVries, A. L. (1986). In “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 293303. Academic Press, Orlando, Florida. Doster, W., Bachleitner, A., Dunau, R., Hiebl, M., and Luescher, E. (1986). Biophys. J. 50, 213-219. Doster, W., Cusack, S., and Petry, W. (1989). Nature (London) 337,754-756. Drapron, R. (1985).N A T O A d v . Study Inst. Ser., Ser. E 90, 171-190. Duckworth, R. B., ed. (1975). “Food Science and Technology Series: Water Relations of Foods.” Academic Press, New York. Durchschlag, H., Puchwein, G., Kratky, O., Schuster, I., and Kirschner, K. (1971).Eur. J. Biochem. 19,9-22. Eden, J., Gascoyne, P. R. C., and Pethig, R. (1980).J. Chem. SOC.,Furuday Trans. I 76, 426-434. Edsall,J. T. (1953). In “The Proteins” (H. Neurath and K. Bailey, eds.), Vol. 1, Part B, lst, pp. 549-726. Academic Press, New York. Edsall,J. T. (1980). ACS Symp. Ser. 127,75-85. Edsall,J. T., and McKenzie, H. A. (1978). Adv. Biophys. 10, 137-207. Edsall,J. T., and McKenzie, H. A. (1983). Adv. Biophys. 16,53-183. Ehrenberg, A., Rigler, R., Graeslund, A,, and Nilsson, L., eds. (1987). Springer Ser. Biophys. 1. Eisenberg, D., and Kauzmann, W. (1969). “The Structure and Properties of Water.” Oxford Univ. Press (Clarendon), London and New York. Eisenberg, D., and McLachlan, A. D. (1986). Nature (London) 319, 199-203. Eisenstadt, M. (1985). Biochemistry 24,3407-3421. Essam, J. W. (1980). Rep. Prog. Pnys. 43,833-912.

PROTEIN HYDRATION AND FUNCTION

165

Feeney, R. E., Burcham, T. S., and Yeh, Y. (1986). Annu. Rev. Biophys. Biophys. C h . 15, 59-78. Fel’dman, Y. D., Valitov, V. M., and Fedotov, V. D. (1986). Stud. Btophys. 111, 111-114. Findsen, E. W., Simons, P., and Ondrias, M. R. (1986). Biochemistry 25,7912-7917. Finney, J. L. (1979). I n “Water: A Comprehensive Treatise” (F. Franks, ed.), Vol. 6, pp. 47-122,411-436. Plenum, New York. Finney, J. L., and Poole, P. L. (1984). C o m m t c Mol. Cell. Btophys. 2, 129- 151. Finney, J. L., Goodfellow,J. M., and Poole, P. L. (1982). NATO A h . Study Zmt. Ser., Sm. A 45,387-424. Finzel, B. C., Weber, P. C., Hardman, K. D., and Salemme, F. R. (1985).J. Mol. Biol. 186, 627-643. Fletcher, P. D. I., Rees, G. D., Robinson, B. H., and Freedman, R. B. (1985). Biochim. Biophys. Acta 832,204-214. Fletcher, P.D. I., Robinson, B. H., and Tabony, J. (1986).J . Chem. Soc., Far&y Tram. Z 84, 231 1-2321. Fletcher, P. D. I., Robinson, B. H., and Tabony,J. (1988).Biochim. Biophys. Acta954,27-36. Flory, P. J. (1941).J. Am. C h . SOC.63, 3091. Flory, P. J. (1953). “Principles of Polymer Chemistry.” Cornell Univ. Press, Ithaca, New York. Foss, J. G., and Reyerson, L. H. (1958).J. Phys. Chem. 62, 1214-1216. Franks, F., ed. (1979). “Water: A Comprehensive Treatise,” Vol. 6. Plenum, New York. Frauenfelder, H., and Gratton, E. (1986). Zn “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 207-216. Academic Press, Orlando, Florida. Frauenfelder, H., Parak, F., and Young, R. D. (1988). Annu. Rev. Btophys. Biophys. C h . 17,451-479. Fucaloro, A. F., and Forster, L. S. (1985). Photochem. Photobiol. 41,91-93. Fujita, Y., and Noda, Y. (1978). Bull. Chem. Soc. Jpn. 51, 1567-1568. Fujita, Y., and Noda, Y. (1979). Bull. Chem. Soc. Jpn. 52,2349-2352. Fujita, Y., and Noda, Y. (1981a). Znt.]. Pept. Protein Res. 18, 12- 17. Fujita, Y.,and Noda, Y. (1981b).Bull. C h m . SOC.Jpn. 54,3233-3234. Fujita, Y., Iwasa, Y., and Noda, Y. (1982).Bull. Chem. Soc.Jpn. 55, 1896-1900. Fullerton, G. D., Ord, V. A., and Cameron, I. L. (1986). Biochim. Biophys. Acta 869, 230-246. Fung, B. M. (1986). In “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 151-161. Academic Press, Orlando, Florida. Gao, J., Kuczera, K., Tidor, B., and Karplus, M. (1989).Scknce 244,1069-1072. Gascoyne, P. R. C., and Pethig, R. (1977).J . Chem. Soc., Far&y Tram. Z 73, 171- 180. Gascoyne, P. R. C., Pethig, R., and Szent-Gyorgyi, A. (1981). Proc. Natl. Acad. Sci. U.S.A. 78,261-265. Gaspar, R., Jr., Andrew, E. R., Bryant, D. J., and Cashell, E. M. (1982). Chem. Phys. Lett. 86,327-330. Gavish, B. (1986). In “The Fluctuating Enzyme” (G. R. Welch, ed.). Wiley, New York. Gavish, B., and Werber, M. M. (1979). Biochemistry 18, 1269- 1275. Gavish, B., Gratton, E., and Hardy, C. J. (1983).Proc. Natl. Acad. Sci. U.S.A. 80, 750-754. Geiger, A., Rahman, A., and Stillinger, F. H. (1979).J. Chem. Phys. 70,263-276. Genzel, L., Kremer, F., Poglitsch, A., and Bechtold, G. (1983).Bzopolymers 22, 1715-1729. Gevorkyan, S. G., and Morozov, V. N. (1983). Biofzika 28,944-948. Giannini, L., and Gratton, E. (1977).Atti Accad. Naz. Lincei, Cl. Sci. Fis., Mat. Nut., Rend. 62, 71 1-715. Gilson, M. K., and Honig, B. H. (1988a).Proteins: Struct., Fund., Genet. 3, 32-52. Gilson, M. K., and Honig, B. (1988b).Proteins: S t r u t . , Funct., Genet. 4,7-18.

166

JOHN A. RUPLEY AND GIORGIO CARER1

Ginzburg, B. Z. (1982).J. Colloid Interface Sci. 85,422-430. Goldanskii, V. I., and Krupyanskii, Y. F. (1989). 4. Rev. Bzqbhys. 22, 39-92. Gonnelli, M., and Strambini, G. B. (1988). J. Phys. Chem. 92,2854-2857. Grant, E. H. (1982). Bioelectrmgnetics 3, 17-24. Grant, E. H., McLean, V. E. R., Nightingale, N. R. V., and Gabriel, C. (1985). NATO Adv. Study Inst. Ser., Ser. A 97,65-74. Gregory, R. B., and Rosenberg, A. (1986). In “Methods in Enzymology” (C. H. W. Hirs and S. N. Timasheff, eds.), Vol. 131, pp. 448-508. Academic Press, Orlando, Florida. Guy, H. R. (1985). Bqfhys.J. 47,61-70. Haggis, G. H. (1956). Biochim. Biophys. Acta 19, 545-546. Haggis, G. H. (1957). Biochim. Biophys. Acta 23,494-503. Hagler, A. T., and Moult, J. (1978). Nature (London) 272,222-226. Haire, R. N., and Hedlund, B. E. (1983). Biochemistry 22,327-334. Halle, B., Anderson, T., Forsen, S., Lindman, B., and Lindman, B. (1981).J . Am. Chem. SOC. 103,500-508. Haly, A. R., and Snaith, J. W. (1968). Biopolymers 6, 1355-1377. Haly, A. R., and Snaith, J. W. (1971). Biopolymers 10, 1681-1699. Hanson, J. C., and Schoenborn, B. P. (1981).J. Mol. Biol. 153, 117-146. Hartmann, H., Steigemann, W., Reuscher, H., and Parak, F. (1987). Eur. Biophys. J . 14, 337-348. Harvey, S. C., and Hoekstra, P. (1972).J. Phys. Chem. 76,2987-2994. Hawkes, J. J., and Pethig, R. (1988). Biochim. Biophys. Acta 952,27-36. Hendrickson, W. A., and Teeter, M. M. (1981). Nature (London) 290, 107- 113. Hermans, J., and Vacatello, M. (1980). ACS Symp. Ser. 127, 199-214. Hill, T. L. (1947).J. Chem. Phys. 15,767-777. Hill, T. L. (1949).J. Chem.Phys. 17, 762-771. Hiltner, A. (1979). Polym. Eng. Sci. 19,722-727. Hilton, B. D., Hsi, E., and Bryant, R. G. (1977).J . Am. Chem. SOC.99,8483-8490. Hirs, C . H. W., and Timasheff, S. N., eds. (1985). “Methods in Enzymology,” Vol. 117. Academic Press, Orlando, Florida. Hnojewyj, W. S. (1971). Proc. N.D. Acad. Sci. 24,84-102. Hnojewyj, W. S. (1978). Proc. N.D. Acad. Sci. 29,67-73. Hnojewyj, W. S. and Reyerson, L. H. (1959).J. Phys. Chem. 63, 1653-1654. Hnojewyj, W. S., and Reyerson, L. H. (1961).J. Phys. C h . 65, 1694- 1698. Holden, H. M., Tronrud, D. E., Monzingo, A. F., Weaver, L. H., and Matthews, B. W. (1987). B i o c h G t r y 26,8542-8553. Honig, B. H., Hubbell, W. L., and Flewelling, R. F. (1986). Annu. Rev. Biophys. Biophys. Chem. 15, 163-193. Hsi, E., and Bryant, R. G. (1975).J. Am. Chem. Soc. 97,3220-3221. Hsi, E., and Bryant, R. G. (1977). Arch. Biochem. Biophys. 183,588-591. Hsi, E., Jentoft, J. E., and Bryant, R. G. (1976a).J. Phys. C h m . SO, 412-416. Hsi, E., Mason, R., and Bryant, R. G. (1976b).J. Phys. Chem. 80,2592-2597. Hutchens, J. O., Cole, A. G., and Stout, J. W. (1969).J. Biol. Chem. 244, 26-32. Hvidt, A,, and Nielsen, S. 0. (1966). Adv. Protein Chem. 21,287-386. Israelachvili, J. N. (1985). “Intermolecular and Surface Forces: With Applications to Colloidal and Biological Systems.” Academic Press, Orlando, Florida. Israelachvili,J. (1987). Acc. Chem. Res. 20,415-421. Israelachvili, J., and Marra, J. (1986). I n “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 353-360. Academic Press, Orlando, Florida. Izumi, T., Yoshimura, Y., and Inoue, H. (1980). Arch. Biochem. Biophys. 200,444-451.

PROTEIN HYDRATION AND FUNCTION

167

Jakobsen, R. J., Wasacz, F. M., Brasch, J. W., and Smith, K. B. (1986). Biopolymers 25, 639-654. James, M. N. G., and Sielecki, A. R. (1983).J . Mol. B i d . 163,299-361. Jorgensen, W. L. (1989).J. Am. Chem. SOC. 111,3770-3771. Kachalova, G. S., Myachin, E. T., Morozov, V. N., Morozova, T. Y., Vagin, A. A., Strokopytov, B. V., and Vol’kenshtein, M. V. (1987). Dokl. Biophys. (Engl. Transl.) 493, 236-238. Kakalis, L. T., and Baianu, I. C. (1988). Arch. Bzochem. Biophys. 267,829-841. Kang, Y. K., Gibson, K. D., Nemethy, G., and Scheraga, H. A. (1988). J. Phys. Chem. 92, 4739-4742. Karle, I. L., and Balaram, P. (1989). UCLA Symp. Mol. Cell. B i d , New Ser. 86, 7 1-85. Karplus, M., and Rossky, P. J. (1980). ACS Symp. Ser. 127, 23-42. Karplus, P. A., and Schulz, G. E. (1987).J.Mol. B i d . 195,701-729. Karplus, M., and Weaver, D. L. (1976). Nature (London) 260,404-406. Kent, M., and Meyer, W. (1984).J.Phys. D 17, 1687-1698. Khurgin, Y. I., Medvedeva, N. V., and Roslyakov, V. Y. (1977). Biofizika 22, 1010-1014. Kim, K., and Kauzmann, W. (1980).J.Phys. Chem. 84,163-165. Kirkpatrick, S . (1979). I n “Matiere Ma1 Condens. [Ec. Ete Phys. Theor.], 31st, Meeting Date 1978” (R. Balian, R. Maynard, and G. Toulouse, eds.), pp. 321-403. NorthHolland, Amsterdam. Klibanov, A. M. (1986). I n “Protein Engineering” (M. Inouye and R. Sarma, eds.), pp. 341 -349. Academic Press, Orlando, Florida. Knight, C. A., and DeVries, A. L. (1989). Science 245,505-507. Koenig, S . H. (1980). ACSSymp. Ser. 127, 157-176. Koenig, S. H., and Schillinger, W. E. (1969).J.B i d . Chem. 244,3283-3289. Koenig, S . H., Hallenga, K., and Shporer, M. (1975). Proc. Nutl. Acad. Sci. U.S.A. 72, 2667-267 1. Kollman, P., Rao, S., Brown, F., Daggett, V., Seibel, G., and Singh, U. C. (1987). UCLA Symp. Mol. Cell. Bwl., New Ser., 69 (Protein S t w t . , FoldingDes. 2), 215-225. Kossiakoff, A. A. (1983). Annu. Rev. Bzophys. Bzoag. 12, 159- 182. Kremer, F., Poglitsch, A., and Genzel, L. (1984). In “Nonlinear Electrodynamic Biological Systems” (W. R. Adey and A. F. Lawrence, eds.), pp. 177-186. Plenum, New York. Krupyanskii, Y. F., Parak, F., Gol’danskii, V. I., Moessbauer, R. L., Gaubman, E. E., Engelmann, H., and Suzdalev, I. P. (1982). Z. Nuturforsch., C: Bzosci. 37C, 57-62. Kundrot, C. E., and Richards, F. M. (1987).J.Mol. Bzol. 193, 157-170. Kundrot, C. E., and Richards, F. M. (1988).J.Mol. Biol. 200,401-410. Kuntz, I. D., Jr. (1971).J.Am. Chem. SOC.93,514-516. Kuntz, I. D., Jr., and Kauzmann, W. (1974). Adv. Protein Chem. 28,239-345. Kurinov, I. V., Krupyanskii, Y. F., Suzdalev, I. P., and Goldanskii, V. I. (1987). Hypofine Interact. 33,223-232. Lee, B., and Richards, F. M. (1971).J . Mol. B i d . 55,379-400. Lee, J. C., Gekko, K., and Timasheff, S. N. (1979). In “Methods in Enzymology” (C. H. W. Hirs and S. N. Timasheff, eds.), VoI. 61, pp. 26-49. Academic Press, New York. Leopold, C., ed. (1986). “Membranes Metabolism and Dry Organisms.” Cornell Univ. Press, Ithaca, New York. Leveque, J. L., Carson, J. C., Pissis, P., and Boudouris, G. (1981). Biopolymers 20, 2649- 2656. Levitt, M., and Sharon, R. (1988). Proc. Natl. Acad. Sci U.S.A. 85,7557-7561. Levitt, M., Sander, C., and Stern, P. S. (1985).J.Mol. B i d . 181,423-447. Likhtenshtein, G. I. (1976). Colloq. Int. C.N.R.S. 246,45-52.

168

JOHN A. RUPLEY AND GIORGIO CARER1

Ling, G. N. (1988). ScanningMicrosc. 2,899-913. Lioutas, T. S., Baianu, I. C., and Steinberg, M. P. (1986). Arch. Biochem. Bzophy. 247, 68-75. Lioutas, T. S., Baianu, I. C., and Steinberg, M. P. (1987).J. A@. Food Chem. 35, 133- 137. Lis, L. J., McAlister, M., Fuller, N., Rand, R. P., and Parsegian, V. A. (1982). Biophys.]. 37, 657-665. Loftfield, R. B., Eigner, E. A,, Pastuszyn, A., Lovgren, T. N. E., and Jakubowski, H. (1980). Proc. Natl. Acad. Sci. U.S.A. 77,3374-3378. Low, P. S., and Somero, G. N. (1975a). Proc. Natl. Acad. Sci. U.S.A. 72,3014-3018. Low, P. S., and Somero, G. N. (1975b). Proc. Natl. Acad. Sci. U.S.A. 72,3305-3309. Luescher, M., and Ruegg, M. (1978). Biochim. Biophys. Acta 533,428-439. Luescher, M., Ruegg, M., and Schindler, P. (1978). Biochim. Btophys. Acta 536,27-37. Luescher, M., Schindler, P., Ruegg, M., and Rottenberg, M. (1979). Biopolj..... 18, 1775-1791. Luescher-Mattli, M., and Ruegg, M. (1982a). Biopolymers 21,403-418. Luescher-Mattli, M., and Ruegg, M. (1982b). Biopolymers 21,419-429. Luisi, P. L., and Steinmann-Hofmann, B. (1987). In “Methods in Enzymology” (K. Mosbach, ed.), Vol. 136, pp. 188-216. Academic Press, New York. Lumry, R., and Gregory, R. B. (1986). In “The Fluctuating Enzyme” (G. R. Welch, ed.). Wiley, New York. Lumry, R., Solbakken, A., Sullivan, J.. and Reyerson, L. H. (1962). J . Am. Chem. SOC.84, 142- 149. Mackay, D. H. J., and Wilson, K. R. (1986).J. Biomol. Struct. Dyn. 4,491-500. Maddox, J. (1988). Nature (London) 332,677. Maroncelli, M., MacInnis, J., and Fleming, G. R. (1989). Science 243, 1674-1681. Martinek, K., Levashov, A. V., Klyachko, N. L., Pantin, V. I., and Berezin, I. V. (1981). Bwchim. Biophys. Acta 657,277-294. Mason, S. A., Bentley, G. A,, and McIntyre, G. J. (1984). Basic Life Sci. 27,323-334. Matthew, J. B. (1985). Annu. Rev. Bzophys. Biophys. Chem. 14,387-417. Matthews, B. W. (1985). In “Methods in Enzymology” (H. W. Wyckoff, C. H. W. Hirs, and S. N. Timasheff, eds.), Vol. 114, pp. 176- 187. Academic Press, New York. McCammon, J. A., Gelin, B. R., Karplus, M., and Wolynes, P. G. (1976). Nature (London) 262,325-326. McLaren, A. D., and Kirwan, J. P. (1976).J. Gen. Virol. 33, Pt. 1, 155-157. McLaren, A. D., and Rowen, J. W. (1951).J. Polym. Sci. 7,289-324. Meyer, E., Cole, G., Radhakrishnan, R., and Epp, 0.(1988). Acta Crystallogr., Sect. B: Struct. Sci. B44,26-38. Mezei, M., and Beveridge, D. L. (1986). Ann. N.Y. Acad. Sci.482, 1-23. Middendorf, H. D. (1984). Annu. Rev. Biophys. Bioag. 13,425-451. Middendorf, H. D., Randall, J. T., and Crespi, H. L. (1984). Basic Life Sci. 27, 381-400. Moras, D., Drenth, J., Strandberg, B., Suck, D., and Wilson, K., eds. (1987). NATOAdv. Study Inst. Ser., Ser.A 126. Morgan, H., and Pethig, R. (1986).J. Chem. SOC.,Faraday Trans. I 82, 143-156. Morozov, V. N., and Gevorkyan, S. G. (1985). Biopolymers 24,1785-1799. Morozov, V. N., Morozova, T. Y., Kachalova, G. S., and Myachin, E. T. (1988). Int. J . Biol. Macromol. 10,329-336. Morozova, T. Y., and Morozov, V. N. (1982).J. Mol. Biol. 157, 173-179. Mrevlishvili, G. M., and Privalov, P. L. (1969). In “Water in Biological Systems” (L. P. Kayushin, ed.), p. 63. Plenum, New York. Nagle, J. F., and Mille, M. (1981).J. Chem. Phys. 74, 1367-1372.

PROTEIN HYDRATION AND FUNCTION

169

Nemethy, G. (1986). In “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 183- 196. Academic Press, Orlando, Florida. Nemethy, G., Peer, W. J., and Scheraga, H. A. (1981). Annu. Rev. Biophys. Bioeng. 10, 459-497. Oefner, C., and Suck, D. (1986).J.Mol. Biol. 192,605-632. Oobatake, M., and Ooi, T. (1988).J. Biochem. (Tokyo) 104,433-439. Ooi, T., and Oobatake, M. (1988a).J. Biochem. (Tokyo) 103, 114- 120. Ooi, T., and Oobatake, M. (1988b). Comments Mol. Cell. Biophys. 5,233-251. Ooi, T., and Oobatake, M. (1988~). J. Biochem. (Tokyo) 104,440-444. Ooi, T., Oobatake, M., Nemethy, G., and Scheraga, H. A. (1987). Proc. Nutl. Acad. Sci. U.S.A. 84,3086-3090. Otting, G., and Wuethrich, K. (1989).J.Am. C h . SOC.111, 1871-1875. Packer, L., ed. (1986). “Methods in Enzymology,” Vol. 127. Academic Press, Orlando, Florida. Parak, F. (1986). I n “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 196-206. Academic Press, Orlando, Florida. Parak, F. (1987). Comments Mol. Cell. Biophys. 4, 265-280. Parak, F. (1989). NATO Adv. Study Inst. Ser., Ser. A 178, 197-221. Parak, F., and Reinisch, L. (1986). In “Methods in Enzymology” (C. H. W. Hirs and S. N. Timasheff, eds.), Vol. 131, pp. 568-607. Academic Press, Orlando, Florida. Parak, F., Hartmann, H., Aumann, K. D., Reuscher, H., Rennekamp, G., Bartunik, H., and Steigemann, W. (1987). Eur. Biophys.J. 15,237-249. Parak, F., Heidemeier, J., and Nienhaus, G. U. (1988). HyperfineInteruct. 40, 147-157. Parsegian, V. A,, Rand, R. P., and Rau, D. C. (1985). Chem. Scr. 25,28-31. Parsegian, V. A., Rand, R. P., Fuller, N. L., and Rau, D. C. (1986). I n “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 400-416. Academic Press, Orlando, Florida. Peemoeller, H., Shenoy, R. K., and Pintar, M. M. (1981).J.Mugn. Resm. 45, 193-204. Peemoeller, H., Kydon, D. W., Sharp, A. R., and Schreiner, L. J. (1984). Cun.J. Phys. 62, 1002- 1009. Peemoeller, H., Yeomans, F. G., Kydon, D. W., and Sharp, A. R. (1986). Biophys. J . 49, 943-948. Permyakov, E. A., and Burstein, E. A. (1977). Stud. Biophys. 64,163-172. Perutz, M. F. (1946). Trans. Furaday SOC.42B, 187- 195. Pessen, H., and Kumosinski, T. F. (1985). I n “Methods in Enzymology” (C. H. W. Hirs and S. N. Timasheff, eds.), Vol. 117, pp. 219-255. Academic Press, Orlando, Florida. Peters, D., and Peters, J. (1986). Biopolymers 25, 1109- 1132. Pethig, R. ( 1979). “Dielectric and Electronic Properties of Biological Materials.” Wdey, Chichester, England. Pethig, R., and Kell, D. B. (1987). Phys. Med. Biol. 32,933-970. Phillips, S . E. V. (1984). Basic Lqe Sci. 27,305-322. Piculell, L., and Halle, B. (1986).J. Chem. Soc., Far&y Trans. 1 82,401-414. Poglitsch, A., Kremer, F., and Genzel, L. (1984).J.Mol. Biol. 173, 137-142. Polnaszek, C. F., and Bryant, R. G. (1984a).J. Am. Chem. SOC.106,428-429. Polnaszek, C . F., and Bryant, R. G. (1984b).J. Chem.Phys. 81,4038-4045. Poole, P. L., and Barlow, D. J. (1986). Biopolymers 25,317-335. Poole, P. L., and Finney, J. L. (1983a). Int. J. Biol. Macromol. 5,308-310. Poole, P. L., and Finney, J. L. (1983b). Biopolymers 22, 255-260. Poole, P. L., and Finney, J. L. (1984). Biopolymm 23, 1647- 1666. Poole, P. L., and Finney, J. L. (1986). I n “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 284-293. Academic Press, Orlando, Florida.

170

JOHN A. RUPLEY AND GIORGIO CARER1

Poole, P. L., Walton, A. R., and Zhang, J. (1987). Int. J . B i d . Macromol. 9, 245-246. Post, C. B., Brooks, B. R., Karplus, M., Dobson, C. M., Artymiuk, P. J., Cheetham, J. C., and Phillips, D. C. (1986).J. Mol. B i d . 190,455-479. Potthast, K., Hamm, R., and Acker, L. (1975). In “Water Related Foods” (R. B. Duckworth, ed.), pp. 365-377. Academic Press, New York. Prabhakaran, M., and Ponnuswamy, P. K. (1980).J. Theor. B i d . 87,623-637. Prouty, M. S., Schechter, A. N., and Parsegian, V. A. (1985).J. Mol. B i d . 184, 517-528. Quiocho, F. A., Wilson, D. K., and Vyas, N. K. (1989). Nature (London) 340,404-407. Radzicka, A., and Wolfenden, R. (1988). Biochemistry 27, 1664- 1670. Rand, R. P., Das, S., and Parsegian, V. A. (1985). Chem. Scr. 25, 15-21. Rao, P. B., and Bryan, W. P. (1978). Biopolymers 17,291-314. Rao, K. S., and Das, B. (1968).J.Phys. C h . 72, 1223-1230. Rau, D. C., Lee, B., and Parsegian, V. A. (1984). Proc. Natl. A d . Sci. U.S.A. 81, 2621-2625. Raymond, J. A,, and DeVries, A. L. (1977). Proc. Natl. Acud. Sci. U.S.A. 74, 2589-2593. Richards, F. M. (1977). Annu. Rev. Biophys. Bioeng. 6, 151-176. Richmond, T. J. (1984).J.Mol. Biol. 178,63-89. Rochester, C. H., and Westerman, A. V. (1976a). J . Chem. SOC., Faruday Trans. 1 72, 2498-2504. Rochester, C. H., and Westerman, A. V. (1976b). J. Chem. SOC., Faraday Trans. 1 72, 2753-2768. Rochester, C. H., and Westerman, A. V. (1977).J . Chem. SOC.,Faruday Trans. 1 73, 33-43. Rosenberg, A. (1986). I n “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 630648. Academic Press, Orlando, Florida. Rosenberg, A,, and Somogyi, B. (1986). Spp.B i d . Hung. 30, 101-112. Roslyakov, V. Y., and Khurgin, Y. I. (1971). Izv. Akad. Nauk SSSR, Ser. Khim., 1366. Rossky, P. J., and Karplus, M. (1979).J.Am. Chem. SOC.101, 1913-1937. Ruegg, M., Moor, U., and Blanc, B. (1975). Eiochim. Biophys. Actu 400,334-342. Ruggiero, J., Sanches, R., Tabak, M., and Nascimento, 0. R. (1986). Can. J. Chem. 64, 366-372. Rupley, J. A., and Siemankowski, L. (1986). In “Membranes Metabolism and Dry Organisms” (C. Leopold, ed.), pp. 259-272. Cornell Univ. Press, Ithaca, New York. Rupley, J. A., Yang, P. H., and Tollin, G. (1980). ACS Symp. Ser. 127, 1 1 1- 132. Rupley, J. A., Gratton, E., and Careri, G. (1983). T r e d B i o c h . Sci. (Pers. Ed.) 8, 18-22. Rupley, J. A,, Siemankowski, L., Careri, G., and Bruni, F. (1988). Proc. Natl. Acud. Sci. U.S.A. 85,9022-9025. Russell, A. J., and Klibanov, A. M. (1988).J. B i d . Chem. 263, 11624- 11626. Saenger, W. (1987). Annu. Rev. Biophys. Biophys. Chem. 16,93- 114. Samanta, S. R., and Walrafen, G. E. (1978).J. C h m . Phys. 68,3313-3315. Savage, H. (1986a). Water Sci. Rev. 2,67-148. Savage, H. (1986b). Biophy5.J. 50,947-965. Savage, H. (1986~). Biophys. J. 50,967-980. Savage, H., and Wlodawer, A. (1986). I n “Methods in Enzymology” (L. Packer, ed.), Vol. 127, pp. 162- 183. Academic Press, Orlando, Florida. Scanlon, W. J., and Eisenberg, D. (1975).J.Mol. B i d . 98,485-502. Scanlon, W. J., and Eisenberg, D. (198 1). J . Phys. Chem. 85,3251 -3256. Schauer, G., Kimmich, R., and Nusser, W. (1988). Biophys.J. 53,397-404. Scher, H., and Zallen, R. (1970).J. Chem. Phys. 53,3759-3761. Scheraga, H. A., and Mandelkern, L. (1953).J.Am. Chem. SOC.75, 179. Schinkel, J. E., Downer, N. W., and Rupley, J. A. (1985). Bzochairtq 24,352-366. Schoenborn, B. P. (1984). Basic Life Scz. 27,261-279.

PROTEIN HYDRATION AND FUNCTION

171

Shablakh, M., Dissado, L. A., and Hill, R. M. (1984).J . B i d . Phys. 12,63-78. Shchegoleva, T. Y. (1984). Biofzika 29,935-939. Sheats, G. F., and Forster, L. S. (1983). Biochem. Biophys. Res. Cummun. 114,901-906. Sherman, F. B., Tusupkaliev, U., Klimova, V. A., and Khurgin, Y. I. (1973). Zzv. Akad. Nauk SSSR, Ser. Khim., 1401 - 1403. Shimanovskii, N. L., Stepanyants, A. U., Lezina, V. P., Ivanov, L. V., and Likhtenshtein, G. I. (1977). Biofizika22, 811-815. Shirley, W. M., and Bryant, R. G. (1982).J. Am. Chem. SOC. 104,2910-2918. Shrake,‘A., and Rupley, J. A. (1973).J. Mol. Biol. 79,351-371. Simatos, D., and Multon, J. L., eds. (1985). NATO Adv. Study Znst. Ser.,Ser E 90. Singh, G. P., Parak, F., Hunklinger, S., and Dransfeld, K. (1981). Phys. Rev. Len. 47, 685-688. Skarzynski, T., Moody, P. C. E., and Wonacott, A. J. (1987).J. Mol. Biol. 193, 171-187. Skujins, J. J., and McLaren, A. D. (1967). Science 158, 1569-1570. Sloan, D. L., and Velick, S. F. (1973).J. Biol. Chem. 248,5419-5423. Sloan, D. L., Samuelson, G. L., Ailion, D. C., and Velick, S. F. (1973). J . Biol. Chem. 248, 5424-5427. Smith, J., Cusack, S., Poole, P., and Finney, J. (1987).J. Biomol. Strzlct. @n. 4,583-588. Smith, J., Kuczera, K., Tidor, B., Doster, W., Cusack, S., and Karplus, M. (1989). Physica B (Amsterdam), 156-157,437-443. Somogyi, B., Norman, J. A., Zempel, L., and Rosenberg, A. (1988). Biophys. Chem. 32, 1-13. Stauffer, D. (1985). “Introduction to Percolation Theory.” Taylor and Francis, Philadelphia, Pennsylvania. Steinhoff, H. J., Lieutenant, K., and Schlitter, J. (1989). Z. Natulforsch., C : Biosci. 44, 280-288. Stevens, E., and Stevens, L. (1979). Biochem. J. 179, 161-167. Stockmayer, W. H. (1943). J. Chem. Phys. 11,45. Stonehouse, J. R. (1982). Dissertation. Wayne State Univ., Detroit, Michigan. Strambini, G. B., and Gabellieri, E. (1984). Photochem. Photobiol. 39,725-729. Strambini, G . B., and Gonnelli, M. (1988).J. Phys. Chem. 92,2850-2853. Suguna, K., Padlan, E. A., Smith, C. W., Carlson, W. D., and Davies, D. R. (1987). Proc. Natl. A c ~S .C ~U.S.A. . 84, 7009-7013. Sundaralingam, M., and Sekharudu, Y.C. (1989). Science 244, 1333-1337. Sundaralingam, M., Sekharudu, Y. C., Yathindra, N., and Ravichandran, V. (1987). Znt.J. Quantum Chem., Quantum Biol. Symp. 14,289-296. Sussman, J. L., Shoham, M., and Harel, M. (1989). Prog. Clin. Biol. Res. 289, 171-187. Suurkuusk, J. (1974). Acta Chem. Scand., Ser. B 28,409-417. Takano, T. (1977a).J. Mol. Biol. 110,569-584. Takano, T. (1977b).J. Mol. Biol. 110,537-568. Tanaka, F., Forster, L. S., Pal, P. K., and Rupley, J. A. (1975). J. B i d . Chem. 250, 6977-6982. Tapia, O., and Eklund, H. (1986). Enzyme 36,101-114. Teeter, M. M. (1984). Proc. Natl. Acud. Sci. U.S.A. 81,6014-6018. Teeter, M. M., and Hope, H. (1986). Ann. N.Y. Acnd. Sci. 482,163-165. Teeter, M. M., and Kossiakoff, A. A. (1984). Basic Life Sci. 27, 335-348. Teeter, M. M., and Whitlow, M. D. (1986). Tram. Am. Crystallogr. Assoc. 22,75-88. Thanki, N., Thornton, J. M., and Goodfellow,J. M. (1988).J. Mol. Biol. 202, 637-657. Tredgold, R. H., Sproule, R. C., and McCanny,J. (1976).J. Chem. Soc., Faraduy Trans. I 72, 509-512. Usha, M. G., and Wittebort, R. J. (1989).J . Mol. Biol. 208,669-678.

172

JOHN A. RUPLEY AND GIORGIO CARER1

Vander Meulen, D. L., and Ressler, N. (1980).Arch. Biochem. Biophys. 199, 197-205. Varo, G., and Keszthelyi, L. (1983). Bzophys. J. 43,47-51. Vedani, A., Huhta, D. W., and Jacober, S. P. (1989). J . Am. Chem. SOC. 111, 4075-4081. Velicelebi, G., and Sturtevant, J. M. (1979). Biochemistry 18, 1180- 1186. Warshel, A., and Russell, S. T. (1984). Q.Rev. Biophys. 17,283-422. Warshel, A., and Sussman, F. (1986). Proc. Natl. Acad. Sci. U.S.A. 83,3806-3810. Warshel, A., and Weiss, R. M. (1980).J. Am. Chem. SOC.102,6218-6226. Warshel, A., Russell, S. T., and Churg, A. K. (1984). Proc. Natl. Acad. Sci. U.S.A. 81, 4785-4789. Warshel, A., Sussman, F., and King, G. (1986). Biochemistry 45,8368-8372. Warshel, A., Sussman, F., and Hwang, J. K. (1988).J. Mol. Bwl. 401, 139-159. Warshel, A., Aqvist, J., and Creighton, S. (1989). Proc. Natl. Acad. Scz. U.S.A. 86, 5820-5824. Watenpaugh, K. D., Margulis, T. N., Sieker, L. C., and Jensen, L. H. (1978).J. Mol. Bzol. 122, 175-190. Watt, 1. C., and D’Arcy, R. L. (1976).J. Polym. Scz., Polym. Symp. 55, 155-166. Weaver, L. H., and Matthews, B. W. (1987).J. Mol. Biol. 193, 189-199. Welch, G. R., ed. (1986). “The Fluctuating Enzyme.” Wiley, New York. Wendoloski, J. J., and Matthew, J. B. (1989). Proteins: S t w t . , Funct., Genet. 5,313-321. Wiggins, P. M. (1988). Prog. Polym. Sci. 13, 1-35. Wiggins, P. M., and MacClement, B. A. E. (1987). Int. Rev. Cytol. 108,249-303. Wilkinson, A., Morowitz, H. J., and Lund, W. (1976).Biophys.J. 16, 193- 197. Winzor, D. J., and Wills, P. R. (1986). Blophys. Chem. 25,243-251. Wlodawer, A,, Svensson, L. A., Sjoelin, L., and Gilliland, G. L. (1988). Biochemistry 27, 2705-27 17. Wodak, S. J., and Janin, J. (1980). Proc. Natl. Acad. Scz. U.S.A. 77, 1736-1740. Wolber, P. K., Deininger, C. A., Southworth, M. W., Vandekerckhove, J., Montagu, M. V., and Warren, G. J. (1986). Proc. Natl. Acad. Sci. U.S.A. 83,7256-7260. Wolfenden, R., Anderson, L., Cullis, P. M., and Southgate, C. C. B. (1981). Biochemistry 20,849-855. Wong, C. H. (1989). Science 244, 1145- 1152. Woodward, C. K., and Hilton, B. D. (1979). Annu. Rev. Btophys. Bzoeng. 8,99- 127. Wright, C. S. (1987).J . Mol. Biol. 194, 501-529. Yagi, T., Tsuda, M., Mori, Y., and lnokuchi, H. (1969).J. Am. Chem.SOC.91,2801. Yang, P., and Rupley, J. A. (1979). Biochemistry 18,2654-2661. Yang, D. S. C., Sax, M., Chakrabartty, A., and Hew, C. L. (1988). Nature (London) 333, 232-237. Young, P. R., and Jencks, W. P. (1977).J. Am. Chem. SOC.99,8238-8248. Zaks, A., and Klibanov, A. M. (1984). Sciace 224, 1249-1251. Zaks, A., and Klibanov, A. M. (1986).J. Am. Chem. SOC. 108,2767-2768. Zaks, A., and Klibanov, A. M. (1988a).J. Biol. Chem. 263,3194-3201. Zaks, A., and Klibanov, A. M. (198813).J. Biol. C h a . 263,8017-8021. Zallen, R. (1979). Stud. Stat. Mech. 7 , 177-228. Zallen, R. (1983). “The Physics of Amorphous Solids.” Wiley, New York. Zallen, R., and Scher, H. (1971). Phys. Rev. B 4,4471-4479. Zientara, G. P., Nagy, J. A., and Freed, J. H. (1980).J. C h a . Phys. 73,5092-5106. Zimmerberg, J., and Parsegian, V. A. (1987).J . Bioenerg. Biomemh. 19, 351-358.

LYSOZYME AND a-LACTALBUMIN: STRUCTURE. FUNCTION. AND INTERRELATIONSHIPS

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By HUGH A McKENZIE* and FREDERICK H WHITE. JR.t 'Department of Chemistry. University College. University of New South Wales. Australian Defence Force Academy. Canberra. ACT 2600. Australia t Department of Chemistry. Florida state Unhrersity. Tallahassee. Florida 32306

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. Early History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Lysozyme and Its Function . . . . . . . . . . . . . . . . . . . . . B. a-Lactalbumin and Its Function . . . . . . . . . . . . . . . . . . . C. Sequence Homology between a-Lactalbumin and Lysozyme . . . . . . I11. Some Aspects of the Occurrence. Isolation, and Characterization of Lysozyme and a-Lactalbumin . . . . . . . . . . . . . . . . . . . . . A. Lysozyme: Occurrence, Isolation. and Kinetics of Cell Lysis . . . . . . B. a-Lactalbumin: Occurrence. Isolation. and Determination of Specifier Activity . . . . . . . . . . . . . . . . . . . . . . . . . IV. Three-Dimensional Structure of Lysozyme . . . . . . . . . . . . . . . A . X-Ray Crystal Structure of Domestic Hen Egg-White Lysozyme . . . . B. Mechanism of Cell Lytic Action . . . . . . . . . . . . . . . . . . . C. Structures of Other Lysozymes . . . . . . . . . . . . . . . . . . . D. Water in Lysozyme Crystals . . . . . . . . . . . . . . . . . . . . V. Three-Dimensional Structure of a-Lactalbumin . . . . . . . . . . . . . A . Models for the Three-Dimensional Structure of a-Lactalbumin (Based on Sequence Homology with Lysozyme) . . . . . . . . . . . . . . . . B. X-Ray Crystal Structure of Baboon Milk a-Lactalbumin . . . . . . . . C. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Comparative Binding of Metal Ions in Lysozyme and a-Lactalbumin . . . . A . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Metal Ion Binding to Lysozyme . . . . . . . . . . . . . . . . . . . C. Metal Ion Binding to a-Lactalbumin . . . . . . . . . . . . . . . . D. Structural Changes on Cation Binding by a-Lactalbumin and Their Implications in Lactose Synthase Activity . . . . . . . . . . . . E . Metal Ion Binding in a-Lactalbumin: Implications for Lysozyme . . . . VII . Amino Acid Composition and Sequence Homologies in Lysozyme and a-Lactalbumin . . . . . . . . . . . . . . . . . . . . . A . Amino Acid Compositions . . . . . . . . . . . . . . . . . . . . . B. Sequence Comparisons . . . . . . . . . . . . . . . . . . . . . . C. Summary of Important Features of Comparative Sequences . . . . . . VIII . Galactosyltransferase and the Lactose Synthase System . . . . . . . . . . A . Galactosyltransferases: Occurrence. Function. and Isolation . . . . . . B. Relationships of Structure to Function in Galactosyltransferase . . . . . C. Interactions of Galactosyltransferase and a-Lactalbumin in the Lactose Synthase System . . . . . . . . . . . . . . . . . . . . D. Structural Requirements of Substrate . . . . . . . . . . . . . . . . E. FinalRemarks . . . . . . . . . . . . . . . . . . . . . . . . . . 173 ADVANCES IN

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IX. Some Additional Physical, Chemical, and Biological Comparisons between Lysozyme and a-Lactalbumin . . . . . . . . . . . . . . . A. Spectroscopic Studies . . . . . . . . . . . . . . . . . . . . . . . B. Small-Angle X-Ray Scattering . . . . . . . . . . . . . . . . . . . C. Electron Spin Resonance and Nuclear Magnetic Resonance . . . . . . D. Association and Aggregation . . . . . . . . . . . . . . . . . . . . E. Denaturation and Renaturation . . . . . . . . . . . . . . . . . . . F. Chemical Reactivities . . . . . . . . . . . . . . . . . . . . . . . G. Immunochemical Properties . . . . . . . . . . . . . . . . . . . . H. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . X. Evolutionary Origins of Lysozyme and a-Lactalbumin . . . . . . . . . . A. Introduction: Molecular Clocks and the Fossil Record . . . . . . . . . B. Evolution of Lysozyme and a-Lactalbumin: Divergenceand/orConvergence? . . . . . . . . . . . . . . . . . . C. Are the Functions of Lysozyme and a-Lactalbumin Mutudly Exclusive? . . . . . . . . . . . . . . . . . . . . . . . . XI. Conclusions and the Future . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . . . . . . . . . .

259 259 265 265 267 268 271 272 275 276 276 280 290 293 299 315

I. INTRODUCTION The initiation of a project to study a selected protein structure requires very careful consideration; it is rather like a decision as to which type of new aeroplane to build. The cost in manpower, time, and money is considerable, and if the structure proves to be obdurate, this expenditure shows little return. Lysozyme, which Dr. Poljak had already studied when he joined the Davy Faraday team in 1960, proved to be a fortunate choice. It is the third protein structure to be successfully analysed, and the first enzyme.

T h e above statement was made by the late Sir Lawrence Bragg (1967) at a Royal Society Discussion on the structure and function of lysozyme held at the Royal Institution on February 3, 1966. A 0.6-nm (6 A) resolution Fourier map of domestic hen egg-white lysozyme had been presented by Blake et al. (1962), and one at 0.2-nm (2 A) resolution was published in 1965 (Blake et al., 1965),just 5 years after Poljak initiated the original study. This work resulted not only in a structure for lysozyme itself, but also in specific information on the active site of the enzyme and the mode of catalysis (Johnson and Phillips, 1965; Blake et al., 196713). Important as this was in expanding our knowledge of enzymatic catalysis, it has had farther-reaching effects. At about the same time (1967), evidence began to accumulate that there was structural homology between lysozyme and the milk protein, a-lactalbumin. Furthermore, the functional role of a-lactalbumin in milk was elucidated for the first time. In the ensuing 20 years much more evidence has accumulated on the structural and functional relationships of these two proteins. During

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1965- 1974 there were some dramatic breakthroughs in our knowledge of the structure of these proteins, but the pace slowed later in that decade. This was followed by a period in which much information was accumulated on the evolution of primary structure and of antigenic sites. The resulting debate did not resolve the questions of convergent and divergent evolution. Despite predicted similarities in the structures of a-lactalbumin and lysozyme, technical and other difficulties resulted in little progress on the elucidation of the three-dimensional structure of a-lactalbumin. The discovery of the essential binding of calcium(I1) by a-lactalbumin in 1980 and ensuing studies resulted in a further attack on this obdurate problem. Success was finally achieved in 1986, and the results have far-reaching implications for the structural, evolutionary, and functional relationships of these two proteins. There have been previous reviews on a-lactalbumin and lysozyme, especially during 1970- 1975. The reviews on lysozyme include the conference proceedings edited by Osserman et al. (1974) and the reviews by Imoto et al. (1972), Jolles and Jollb (1984), and Proctor and Cunningham (1988). Work on a-lactalbumin and lactose synthase has been reviewed by Brew (1970), Hill and Brew (1975), Brew and Hill (1975), and Hall and Campbell (1986). In this article we have a somewhat different purpose: to consider both the structure and function of a-lactalbumin and lysozyme in relation to each other, especially in light of the work done in the past few years. We consider also the potential significance of the studies in health and the pathology of disease, such as cancer. We open with a brief report of the early discovery of the occurrence and isolation of these proteins and the elucidation of their function and homology, followed by a brief discussion of some problems in their isolation and the determination of their activity. We then consider various aspects of their three-dimensional structures and their significance. We summarize studies on the implications of their sequence similarities, and also on the binding of metal ions, especially calcium(II), and consider their implications. Then follows a brief discussion of lactose synthase, an enzyme of which a-lactalbumin and galactosyltransferase are essential components. We then examine the evidence concerning the evolution of the two proteins, about which there are conflicting views (see Section X,B). Some conclusions and predictions of future directions are made. The subjects chosen and the emphasis placed on certain aspects reflect the experience and interest of the authors. We make no attempt to be encyclopedic, and the omission of any work does not imply that it is unimportant. However, we do hope that those not working in this field

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will appreciate the progress that has been made and that we also give some inspiration and challenge to others working in this area. 11. EARLYHISTORY A. Lysozyme and Its Function It has been reported that the early Romans used egg white in the treatment of eye infections, and some mothers are reputed to have used human milk for the same purpose; both media contain appreciable amounts of lysozyme. Later, antibacterial properties of leukocytes (Metschnikoff, 1883),cow milk (Fokker, 1890), Bacillussubtilis (Nicolle, 1907), domestic hen egg-white (Laschtschenko, 1909), and nasal secretions (Bloomfield, 1919, 1920) were described from 1880 to 1920. It has been stated (e.g., Fleming and Allison, 1922) that none of the workers prior to 1920 considered “bacteriolysis” in their publications. Nevertheless, Nicolle ( 1907) does discuss “taction bacttkiolysente” in his paper. While Laschtschenko (1909) does not consider bacteriolysis, he does attribute the spore-destroying properties of egg white to “proteolytic enzymes.” However, it was Fleming who first clearly showed that an enzymic substance present in a wide variety of secretions is capable of rapidly lysing (i.e., dissolving)certain bacteria, particularly a yellow “coccus”that he studied. Fleming’s chief, Almroth Wright (quoted by Maurois, 1959), who delighted in constructing words from Greek roots, suggested that the substance be called lysozyme and that the microbe be called Micrococcus lysodeikticus. In December 1921 Fleming presented a paper on his work to the Medical Research Club in London, but it was coldly received. However, the icy reception did not deter Fleming (1922) from submitting his classical paper, “On a Remarkable Bacteriolytic Element Found in Tissues and Secretions,” to the Royal Society in February 1922. Despite the lack of interest, Fleming continued his work on lysozyme for several years and remained convinced of its importance. In 1936 Meyer et al. showed that lysozyme is a protein, and in the following year Abraham and Robinson (1937) first reported its crystallization. Various biochemical and chemical investigations were made of lysozyme over the next 30 years, but it was not until the 1960s that Fleming’s great faith in lysozyme was vindicated. Although Fleming showed the presence of lysozyme in an amazing variety of secretions, he does not appear to have been the first to show its presence in milk. Bordet and Bordet (1924) showed that it is present in human colostrum and milk, but failed to detect it in cow milk. Later, Fleming (1932) concluded that it was present in cow milk, but at a much

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lower level than in human milk. Indeed, the level of lysozyme in cow milk became controversial, and some workers even disputed its presence at all. This controversy has only recently been resolved (see Section 111,A). The nature of the role of lysozyme in the lysis of bacteria was finally elucidated by Salton and co-workers nearly 40 years after Fleming’s original work (Salton, 1964). It had been evident from the earlier studies by Meyer et aZ. (1936) and Epstein and Chain (1940) that the characterization of the substrate for lysozyme and the products formed as a result of its action should throw light on the nature of the structure of the bacterial cell wall. Salton (1952) subsequently showed that the isolated cell wall of M. Zysodeikticus could be used as the substrate for lysozyme and that this could be exploited to determine the nature of the digestion products. A series of studies showed that the lysis of bacteria by lysozyme involved a specific cleavage of cell wall mucopolysaccharides. Of the saccharides liberated, the simplest found was a disaccharide; its structure was investigated by Perkins (1960) and by Salton and Ghuysen (1959, 1960). Since their original work there has been some refinement in our knowledge of the nature of the linkage cleaved between N-acetylglucosamine and N-acetylmuramic acid [/3( 1 4 4 ) linkage, not /3( 1-6) as originally proposed]. A schematic diagram of the cleavage is shown in Fig. 1.

---

o-(NAM)--CH3CHCOOH

I

(NAM)

I

(NAG)

Lysozyme

(NAM)

(NAG)

FIG. 1. Catalysis by lysozyme (1,4-p-N-acetylmuramidase)of the cleavage of the glycosidic linkage between the C-1 of N-acetylmuramic acid (NAM) and the C-4 of Nacetylglucosamine (NAG) in a polymer of NAM and NAG. The vertical broken line shows the point of cleavage.

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HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

In this period it was also shown by Berger and Weiser (1957) that lysozyme can also degrade highly purified chitin, and it was proposed that lysozyme and related enzymes be called muramidases (Salton, 1964). Lysozymes are now defined as 1,4-P-N-acetylmuramidases cleaving the glycosidic linkage between the C- 1 of N-acetylmuramic acid and the C-4 of N-acetylglucosamine in the bacterial peptidoglycan. B . a-Luctulbumin and Its Function

From 1885 to 1965 work on another protein, a-lactalbumin, was proceeding independently of, and without apparent relevance to, that on lysozyme. It is difficult to assess some of the early work because of confusion in nomenclature, especially with respect to the terms “lactalbumin” and “lactoglobulin” (McKenzie, 1967). The first extensively reported study of lactalbumin in bovine milk and colostrum seems to be that by Sebelien (1885), who also referred to some earlier analytical studies of lactalbumin. However, the first reported crystallization of a lactalbumin is that by Wichmann (1899), who appears to have established that his preparation is different from serum albumin (see also Mann, 1906). Following Svedberg’s development of the ultracentrifuge, it was possible in the 1930s to perform ultracentrifugal studies of skim milk and proteins isolated by this method. The noncasein fraction was found to exhibit three peaks, designated as a, P, and 7 , in sedimentation velocity patterns (Sjogren and Svedberg, 1930).The sedimentation coefficient of the /3 peak was shown by both Phillipi and Pedersen in 1935-1936 to be identical to that of the lactoglobulin isolated by Palmer (see Pedersen, 1936a). In 1935 Kekwick isolated a lactalbumin from milk, and soon afterward Pedersen (1936b) concluded from ultracentrifugal studies that it was the protein responsible for the a peak. [It was later shown to be similar to the “crystalline insoluble substance” of Sorensen and Sorensen (1939).] Thus, the two proteins were called P-lactoglobulin and a-lactalbumin. Although it was soon established that these two proteins were the dominant “whey” proteins of cow milk, their functions proved to be elusive. Indeed, the biological function of the former is still uncertain, and it was only 20 years ago that the function of the latter was established. In the early 1960s Hassid and collaborators demonstrated that the enzyme lactose synthetase (now synthase) exists as a microsomal enzyme in the mammary glands of lactating cows and guinea pigs (Watkins and Hassid, 1962) and in a soluble form in cow milk (Babad and Hassid, 1964, 1966). They confirmed an earlier suggestion by Wood and co-

179

LYSOZYME AND a-LACTALBUMIN

workers that lactose is enzymatically synthesized in the mammary gland from UDP-galactose and glucose (see Brew and Hill, 1975). Soon afterward, Brodbeck and Ebner (1966) showed that the milk enzyme (or the microsomal enzyme when solubilized by sonic oscillation) could be resolved into two protein fractions, A and B. Neither A protein nor B protein alone exhibited lactose synthase activity, but recombination of the two fractions restored catalytic activity. Subsequently, Ebner et al. (1966; see also Brodbeck et al., 1967) showed that the B protein is a-lactalbumin. The substrate specificity of the lactose synthase system was studied further by Brew et al. (1968). They confirmed that neither A protein nor B protein alone was active for the synthesis of lactose, but the A protein catalyzed the following reaction (see also Fig. 2): UDP-galactose

+ N-acetylglucosamine + N-acetyllactosamine

+ UDP

(1)

Thus, protein A is a galactosyltransferase (UDP-galactose :N-acetylglucosamine-P-4-galactosyltransferase;EC 2.4.1.38). Its normal function

UDP-Gal

NAG

NAL (a)

UDP-Gal

Glucose

Lactose

(b) FIG.2. Reactions catalyzed by galactosyltransferase (GT). (a) The incorporation of galactose (Gal) into a p(1-4) linkage with N-acetylglucosamine (NAG) to form N-acetyllactosamine (NAL). UDP, Uridine diphosphate. (b) Modification of the activity of GT by a-lactalbumin (a-LA) to convert it to a lactose synthase catalyzing the formation of lactose from UDP-Gal and glucose.

180

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

is to catalyze the incorporation of galactose (Gal) into p( 1+4) linkage with N-acetylglucosamine (N-AcGlc) during the synthesis of the oligosaccharide prosthetic groups of certain glycoproteins (Hill et al., 1969): UDP-Gal

+ N-AcGlc + Gal-N-AcGlc + UDP

1 I Protein

(2)

Protein

With the onset of lactation, a-lactalbumin is formed in the mammary gland and alters the substrate specificity of the transferase from N acetylglucosamine to glucose, enabling lactose synthesis to be effected: UDP-Gal

+ glucose + lactose + UDP

(3)

[Note that the protein glycosylation reaction (2) is not inhibited when a-lactalbumin is formed.] Hereafter we will not use the terms “A protein” and “B protein,” but will refer to them as galactosyltransferase and a-lactalbumin, respectively. Because of its unique ability to alter the specificity of the galactosyltransferase to a lactose synthase (UDP-D-galactose:D-glucose P-~-Dgalactosyltransferase; EC 2.4.1.22), a-lactalbumin has been designated a “specifier” protein. A unique biological role (other than a minor nutritional one) finally had been demonstrated for a-lactalbumin. Although there are several enzymes that consist of two proteins, the unique feature of the involvement of a-lactalbumin is that it is able to change the acceptor specificity of the galactosyltransferase (Ebner, 1970).

C. Sequence Homology between a-Lactalbumin and Lysozyme In 1958 Yasunobu and Wilcox drew attention to certain similarities between a-lactalbumin and lysozyme (see Gordon, 1971). A few years later Brew and Campbell (1967) also drew attention to their marked similarity in molecular weights, amino acid composition, and the aminoand carboxy-terminal amino acid residues. They stated, “To the extent that the properties mentioned reflect similar primary structures, the a-lactalbumins may have evolved by gradual modification from lysozyme, which is found in the milk of many species” (p. 263). This proposal prompted Brew et al. (1967, 1970) to determine the amino acid sequence of bovine a-lactalbumin, which proved to have a high level of sequence identity with domestic hen egg-white lysozyme. Thus, these studies were in accordance with the proposal that the two proteins had diverged from a common ancestor (see also Hill et al., 1969, 1974). They stated that “although lysozyme does not participate in lactose synthesis and a-lactalbu-

LYSOZYME AND a-LACTALBUMIN

181

min does not act on lysozyme substrates, the integral role of a-lactalbumin in lactose synthesis implies a functional as well as a structural similarity between a-lactalbumin and lysozyme. One enzyme is involved in the cleavage and the other in the synthesis of a p( 1+4)-glucopyranosyl linkage.’’ We will discuss in Section X,C how far these statements need modification in light of subsequent work. 111. SOMEASPECTS OF THE OCCURRENCE, ISOLATION, AND CHARACTERIZATION OF LYSOZYME AND ~-LACTALBUMIN A. Lysozyme: Occurrence, Isolation, and Kinetics of Cell LyszS

Lysozyme occurs in domestic hen egg-white to the extent of -30 mg g- ’. It is the most extensively studied lysozyme and is representative of a class of lysozymes, designated chicken- or chick-type lysozymes, now usually abbreviated c-type lysozymes. Although the majority of amino acid sequences of egg-white lysozymes determined have been for the c type, this type has been found at high concentration in only two orders of birds: the Galliformes and the Anseriformes. The c-type lysozymes consist of a single amino acid chain of 129 residues and a molecular weight of -14,500. A different type of lysozyme was found by Dianoux and Jollks (1967) and by Canfield and McMurry (1967) in Embden goose egg white. It was shown by Prager et al. (1974) to be present in at least nine orders of birds. This lysozyme has been designated to be of the goose type, usually abbreviated g type. In the egg white of the black swan (Cygnus atratus) both c and g types occur. Despite their ubiquity, few sequences of the g type have ever been determined. Each contains 185 residues, with an M , of -20,500 (Canfield et al., 1971; Simpson et al., 1980; Simpson and Morgan, 1983). All mammalian lysozymes thus far examined have proved to be of the c type. Bacteriophage lysozymes contain about 164 amino acid residues with M , 18,700. Fungus and bacterial lysozymes show considerable differences from the c type (Fouche and Hash, 1978). Powning and Davidson (1976) have characterized a c-type lysozyme from the wax moth (Galleria mellonella). Jolles et al. (1979b) found the c type among some members of the insect order Lepidoptera, but found a different type in eggs of Ceratitis capitata. By DNA sequencing, Engstrom et al. (1985) unequivocally demonstrated that the Hyalophoru (moth) lysozyme is c type. Other invertebrate lysozymes have been discussed by Jollks and Jollks ( 1984). T h e first plant lysozyme appears to have been isolated from papaya latex by Smith et al. (1955).

-

-

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HUGH A. MCKENZIE A N D FREDERICK H . WHITE, JR.

Because of the high isoelectric point and limited molecular size, it is not surprising that isolation of the lysozymes has proceeded generally along fairly similar paths. In the isolation from milk and egg white from a variety of species, the lysozyme has been adsorbed on an ion exchanger, such as Amberlite IRC-50 (Rohm and Haas, Philadelphia, PA), CM-cellulose (Whatman Inc., Clifton, NJ), CM-Sephadex (Pharmacia, Uppsala, Sweden), and BioGel-CM 30 (Bio-Rad, Richmond, CA), from an ammonium sulfate fraction of the original egg white, or skim milk. After elution it is often purified further by gel filtration on Sephadex G50 or another suitable medium. Whitaker (1963) noted abnormal retention on Sephadex gel under certain conditions. He attributed this to interaction with the dextran, possibly related to the enzymatic function. In more recent work selective removal from immunoadsorbents (e.g., MacKay et al., 1982), heparin-Sepharose (e.g., Boesman-Finkelstein and Finkelstein, 1982; Teahan et al., 1991b), and chitin (e.g., Grinde et al., 1988) was effected. Isoelectric focusing (e.g., Lie and Syed, 1986) has also been used. There are several problems requiring careful attention. Lysozyme has a tendency to form complexes with many substances [e.g., alkyl sulfates, fatty acids, aliphatic alcohols (Smith and Stocker, 1949), cephalins (Brusca and Patrono, 1960), and other proteins]. Of particular importance is its tendency to form complexes with transferrins [e.g., ovotransferrin (Ehrenpreis and Warner, 1956)l. These interactions lead to difficulties in the isolation of lysozyme. Some recent workers have used fast protein liquid chromatography (FPLC) and high-performance liquid chromatography (HPLC) (e.g., Ekstrand and Bjorck, 1986). The resolution in these procedures may not always be satisfactory, and in HPLC pressure and solvent effects must be monitored carefully if the product is to be suitable for conformation and activity studies. There have been special problems in the isolation of some lysozymes, particularly those from cow milk and the milk of monotremes. Isolation from colostrum has proved difficult because of the previously noted tendency of lysozyme to complex with other proteins (e.g., immunoglobulins). Some proteins, particularly lactoferrin and transferrin, may elute from ion exchangers similarly to lysozyme, and it is frequently necessary to take the precaution of using multiple passages of the lysozyme-containing product thereafter, through an appropriate gel filtration column. The isolation of lysozyme from cow milk proved especially difficult. There were three problems: (1) Lysozyme is present in low concentration (-100 pg liter1), necessitating a large volume of starting milk. (2) The enzyme is unstable in raw milk, requiring the immediate start of the processing procedure (White et al., 1988), after which the enzyme is

LYSOZYME A N D (Y-LACTALBUMIN

183

more stable. The instability in fresh milk may be due to proteolytic action on the lysozyme. (3) The enzyme was found to be closely associated with another protein in the final stage of purification, from which lysozyme could be separated only partially, but sufficiently to permit partial amino acid sequence determination (White et al., 1988). This problem of separation had not been realized earlier, when Chandan et al. (1965) reported isolation of cow milk lysozyme, and the subsequent characterization of their product was unknowingly done on a mixture of lysozyme and the contaminating protein. This combination of proteins which were difficult to separate was first observed in attempts to undertake sequencing of the milk lysozyme that had been purified by White et al. (1988). The results showed the existence of two molecular species, one minor, amounting to -30% by weight. This species, however, was identifiable as a c-type lysozyme by its partial sequence. The partial sequence of the remaining component was not identifiable with any known protein sequence. A partial separation could be obtained with HPLC, eluting with dilute acetic acid. T h e trailing edge of the more slowly moving component was confirmed as lysozyme by activity determination and partial sequencing to demonstrate the presence of lysozyme. The question can now be raised as to whether other lysozymes “purified” from sources of low lysozyme levels might be similarly contaminated. Most of the methods by which lysozyme activity has been detected and studied are variations on the principle first discovered by Fleming, that is, the lysis of bacterial cell walls. Typically, a turbid cell suspension (Mzcrococcus luteus) is observed to clear in the presence of lysozyme, and the rate of change of turbidity is equated with lysozyme activity. Some recent workers (e.g., Hammer and Wilson, 1990) detect lysozyme in electrophoretic gels by overlaying the gel with another gel containing M. luteus cells [see also Osserman and Lawlor (1966) and the modification by Lie et al. (1986)l. Also noteworthy as a means for lysozyme determination is radioimmunoassay (Canfield et al., 1974). Another means involves use of the synthetic substrate 3,4-dinitrophenyltetra-N-acetyl-P-chitotetraoside with a spectrophotometric determination (Ballardie and Capon, 1972). This substrate, however, did not lend itself to high sensitivity determinations and was not found to be satisfactory for the determination of lysozyme at low concentration levels (our unpublished observations). A highly sensitive, yet practical, means of lysozyme determination is that of McKenzie and White (1986), which uses the turbidimetric principle. An important feature of their method is the prolonged incubation of the reaction mixture so as to magnify traces of enzymatic activity.

184

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

Although extended reaction time was also used by Selsted and Martinez (1980) in their turbidimetric method, they did not stress the importance of the kinetic order of cell lysis. McKenzie and White (1986) showed that, while the reaction is biphasic, simple kinetics are obeyed over a sufficiently long period that this property can be exploited in the determination. The kinetic order must be considered in the quantitative treatment of the results. The limit of detection was found to be 100 pg ml-' of reaction mixture, equivalent in the case of bovine milk lysozyme to 6 ng ml-' of skim milk, for a 50-1.11 sample. The method of McKenzie and White (1986) readily lends itself to the study of reaction kinetics. A number of lysozymes have been assessed according to kinetic order in the primary phase. Thus, Table I shows that second order predominates in either Tris or imidazole buffer, except that zero order is observed for the platypus milk and tammar wallaby stomach mucosal enzymes. A range of kinetic orders is observed in phosphate buffer, with equine and domestic hen egg-white lysozymes being first order, and others being either second or zero order. Particularly difficult to understand is the exhibition of second-order kinetics for a reaction that basically involves a single reactant, that is, the TABLE I Kinetic Ordersfor Lysis of Cells of Micrococcus l&us with Lysozymes from Various Sources"

Kinetic order Enzyme source Domestic hen egg white Black swan egg white, c and g Human milk Horse milk Cat milk' Echidna milk Platypus milk Cow milk Bovine stomach mucosa Tammar wallaby stomach mucosa

Imidazole or Trisb Phosphateb 2 2 2 2 2 2

0

0

0

2 2

2

0

2 2 1

2 2 1 0

"Different buffer systems for reaction. All results are from McKenzie and White (1986, 1987), except where indicated. Reaction mixture buffer. 'Unpublished results of J. Halliday, H. A. McKenzie, and F. H. White, Jr.

LYSOZYME AND Q-LACTALBUMIN

185

bacterial cell wall. Various workers have reported kinetics of cell lysis with lysozyme as being zero order (Shugar, 1952; Smolelis and Hartsell, 1949), first order (Dickman and Proctor, 1952; Kerby and Eadie, 1953), or second order (Smith et al., 1955; Howard and Glazer, 1969; Prasad and Litwack, 1963). The latter group speculated that the phenomenon of second order in the main phase might be caused by two points of attack on the cell surface. They suggested that lysozyme may not release readily from the cellular debris after cell wall cleavage and that the ensuing slow stage involves cleavage of one linkage at a time. Howard and Glazer (1969) later presented a detailed sequential mechanism for the mode of action of papaya lysozyme. Another possibility for explaining second-order kinetics is that each lysozyme molecule, being trapped in the cellular debris, is, in effect, eliminated from the reaction mixture after it participates in a single lysis. The lysozyme so involved then behaves kinetically as if it were a reactant rather than a catalyst, since it cannot separate from the product in order to participate in another reaction. However, the extent to which lysozyme behaves in this way, as a second reagent, to account for secondorder kinetics, remains to be assessed. M. F. Hammer and A. C. Wilson developed a method by which lysozyme can be detected after the electrophoresis of biological fluids (Dobson et al., 1984). In this method the unstained electrophoretic gel was overlaid with a polyacrylamide gel containing a suspension of M. luteus cells. The lysozyme was allowed to diffuse into the overlay gel, where it catalyzed lysis of the suspended cells. The resulting change in opacity of the overlay could then be quantified by densitometric scanning. In a recent modification of this method (Cortopassiand Wilson, 1989), the sensitivity of lysozyme detection was increased 25-fold, being in the nanogram range. An additional means of quantifying the lysozyme after electrophoresis, again involving an overlay containing a suspension of M. luteus cells, exploited a different, previously untried, principle. Advantage was taken of the fact that lysis of the bacteria cells releases a number of dehydrogenases, principal among which is isocitrate dehydrogenase. The staining method used is suitable for all enzymes that regenerate NADPH or NADH and involves the formation of an insoluble tetrazolium dye (Giblett, 1969). Thus, staining of the overlay reveals the position and intensity of the lysozyme, which can be quantified by densitometric scanning. Moser et al. (1988) described a fluorometric method of detection of lysozyme down to levels of 0.1 pg. They used highly purified cell walls to prepare a peptidoglycan which is labeled with fluorescamine. After

186

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

hydrolysis of a suspension of the fluorescamine-labeled proteoglycan for a fixed time, followed by filtration, the fluorescence of the filtrate is determined and the lysozyme level is estimated. Morsky (1988) has described a method for detecting human lysozyme (with a sensitivity of 5 5 ng) in body fluids, by a procedure involving immunoblotting.

B . a-Lactalbumin: Occurrence, Isolation, and Determination of Specifier Activity Although lysozymes have been isolated from milk, tears, egg white, plant and insect materials, etc., a-lactalbumin has been found only in milk and colostrum (Table 11). However, there is now some evidence for a-lactalbumin-like proteins occurring elsewhere (e.g., in the male reproductive tract). Until the mid-l970s, there was little investigation of the proteins of milk in species other than bovine. A solid body of knowledge of the cow milk proteins has been built up gradually, so that they have become a valuable point of reference for the proteins of other mammals. During the past 10 years sufficient evidence has accumulated to indicate that there are both significant qualitative and quantitative differences in the protein composition of the other species. For example, the casein group of proteins is the dominant protein group in cow milk, but in human milk the whey proteins (i.e., noncasein proteins) quantitatively exceed the caseins. The major whey protein in cow milk is p-lactoglobulin, present in mature phase milk of Western dairy breeds, to the extent of 2.6-4.0 g 1- l , followed by a-lactalbumin at 1 g I-' (McLean et al., 1984). In contrast, human milk contains, at most, traces of plactoglobulin, possibly none of which is synthesized de nova in the human mammary gland. a-Lactalbumin is the dominant whey protein. The occurrence of any a-lactalbumin in the milk of the monotreme Tachyglossus aculeatw multiaculeatw is controversial. Thus, the level of a-lactalbumin in the milk of different species can vary from trace levels (or even zero) to concentrations on the order of 1 g I-'. Because of this wide range in concentration, the complex protein composition of milk, and the even greater complexity of colostrum, considerable care must be exercised in the isolation of a-lactalbumin (indeed, the same may be said of the isolation of lysozyme from these secretions). Furthermore, if the study of the isolated protein is to include conformation and/or studies of enzymatic (and immunological) activity, considerable care must be taken that the method of isolation does not alter the conformation or interfere with the activity. This is particularly important in determining whether or not the isolated protein has a-lactalbu-

-

LYSOZYME AND a-LACTALBUMIN

187

min-like or lysozyme-like activity, or a major activity of one type and a minor activity of the other. In dealing with the milk of rare monotremes and marsupials, samples of which are limited in volume, fractionation by HPLC methods might appear very attractive. However, because of the pressures involved and the nature of the solvents frequently used, both conformation changes and loss of activity may occur. Thus, considerable care should be exercised in the use of HPLC. While known sequencing methods may enable the sequence o r partial sequence to be established for an impure protein [e.g., cow milk lysozyme (White et al., 1988)], protein of high purity is required for other purposes (e.g., determination of trace lytic activity in an isolated alactalbumin or trace lactose synthase specifier activity in a lysozyme). Otherwise, erroneous conclusions may be drawn with respect to structure and function and their evolutionary relationships. Even for the well-characterized cow milk, considerable care must be exercised in the isolation of a-lactalbumin. Armstrong et al. (1967, 1970) made a careful study of the salt-pH solubility relationships of cow “whey” obtained by fractional ammonium sulfate precipitation from whole milk. If the pH is too low (i.e., 5 3 ) , irreversible changes may occur in the state of association and of conformation of both a-lactalbumin and P-lactoglobulin. Although there is evidence (Kronman et al., 1965) that a-lactalbumin, for example, undergoes reversible changes at low pH, it is the experience of one of the authors (H. McK.) that the behavior of the pure individual proteins is not necessarily always the same as that of the respective protein of the a-lactalbumin-P-lactoglobulin mixture in the “whey” medium. On the other hand, there are constraints on the higher pH side as well: The pH must be sufficiently high to maintain a reasonable solubility of the a-lactalbumin, but not so high that it undergoes aggregation (P-lactoglobulin is also sensitive to pH >7.0). It is evident from recent studies of metal ion binding to a-lactalbumin that spurious results will be obtained if aggregation occurs and if the order of mixing is varied. Diuturnal effects have been observed. Metal ion binding studies are critical to our future understanding of the mechanism of action of a-lactalbumin. Thus, it is critical that reaction only be performed with freshly prepared a-lactalbumin that has not been subjected to conditions that may cause irreversible or quasiirreversible changes. Thus, it is best, if possible, to work in a narrow pH region during the fractionation. It is generally possible in the column chromatography of whey proteins to achieve satisfactory fractionation between pH 6.3 and 7.5. Lindahl and Vogel (1984) studied the purification of bovine, human, caprine, ovine, and equine a-lactalbumins, exploiting the property of

TABLE I1 a-Lactalbumin in Milk of Various Specks Species Bovine (Bos)

Variant

B: Most common variant, occurs in Bos taurus, Bos indicur, Bos (Poephagw g r u n n i m ) ; A: Occurs

Note

Refs."

B and A variants, homozygotes shown to contain 1 major, 3 minor components

1

Major and possible minor components Major and possible minor components Heterogeneity observed in camel Similar to, but not identical with, baboon (Papio cynocephalw) and chimpanzee (Pan troglodytes) Minor component also in each homozygote B and C variants isolated from colostrum of Arabian horse (Equus caballus caballus perissodactyla)

2

in Bos indicw, Bos taurus, and crosses; C: Occurs in Bos (bzbos) javanicus w

00 00

Sheep (OvW aries) Goat (Capra hircus) Camel (Camelus dromedarius) Human (Homo sapienr)

Pig (Sw scrofa) Horse (Equus caballus)

Guinea pig (Cavia porcellus) Rabbit (Oryctolagw cuniculw) Rat (Rattwnoruegzcus)

B: Most common variant; A: Less common A: Common variant

Glycoprotein 140 residues (chain extension and is glycoprotein)

3 4

5

6

7

8 9 10

I

02 rc)

Mouse ( M u musculus) Dog (Canisfamiliaris) Cat (FelW catus) Marsupials Red kangaroo (Macropus rufus) Grey kangaroo (Macropus giganteus) Tammar wallaby (Macropus eugenii) Red-necked wallaby (Macropzcs rufognseus) Ring-tailed possum (Pseudocheirtu peregnnus) Monotremes Echidna (Tachyglossus aculeatus) Platypus (Ornithorhynchw anatinus)

11 12 13 14 Two variants

One variant present throughout lactation

15 16 17 18

? ?

Occurrence controversial

19

20

“References: (1) Aschaffenburg and Drewry (1957), Bell et al. (1970, 1981a), Blumberg and Tombs (1958), Gordon (1971), Grosclaude et al. (1976), Hopper and McKenzie (1973), Proctor et al. (1974); (2) Bell and McKenzie (1964), Schmidt and Ebner (1972); (3) Jenness (1982), Schmidt and Ebner (1972); (4) Beg et al. (1985), Conti et al. (1985); (5) Findlay and Brew (1972), Hanson (1960),Jenness (1982), Nagasawa et al. (1973), Phillips and Jenness (1971), R. Greenberg, unpublished observations (see Stuart et al., 1986); (6) Bell et al. (1981~); (7) Bell et al. (1981b), GodovacZimmermann et al. (1987); (8)Brew and Campbell (1967), Brew (1972); (9) Hopp and Woods (1979),Quarfoth and Jenness (1975); (10) Brown et al. (1977), Nicholas etal. (1981), Prasad et al. (1982), Qasba and Chakrabartty (1978); (11) Nagamatsu and Oka (1980), Bhattacharjee and Vonderhaar (1983); (12) Quarfoth and Jenness (1975); (13) Halliday etal. (1990); (14) Bell et al. (1980), McKenzie etal. (1983); (15) Bell et al. (1980), Brew et al. (1973); (16) Nicholas etal. (1987); (17) Shewale etal. (1984); (18) Nicholas etal. (1989); (19) Hopper and McKenzie (1974), Teahan (1986), Teahan etal. (1991a); (20) Teahan et al. (1991b).

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HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

their binding to phenyl-Sepharose in the presence of EDTA and their elution in the presence of Ca(I1). Nevertheless, they caution against using this procedure to determine the quantitative binding of metal ions to a-lactalbumin, because the hydrophobic support stabilizes one conformation, and the binding constants are then not comparable to those determined in free solution. It is now well known that a-lactalbumin may exhibit apparent as well as true heterogeneity. This apparent heterogeneity exhibited in electrophoresis and column chromatography probably involves interactions with buffer ions, but appears to be complex in nature (see, e.g., Gordon, 1971; Hopper, 1973; Prieels and Schlusselberg, 1977). In view of the recent demonstration that a-lactalbumin is a metalloprotein (containing calcium), and the finding by Fenna (1982a) that a-lactalbumin only gave apparent heterogeneity on columns when not fully saturated with Ca(II), it may be that the previous observations on apparent heterogeneity are related, at least in part, to this phenomenon. The occurrence of genetic variants of bovine a-lactalbumin, reflecting autosomal alleles without dominance, is now well established (Table 11). Porcine a-lactalbumin appears to be the only other a-lactalbumin for which similar genetic variation has been established (for references, see Table 11, footnote a). Hopper and McKenzie (1973) observed that each homozygote consists of a major component and three minor ones: one moving faster than the main component and two moving more slowly in electrophoresis at alkaline pH. The fast-moving minor component (F) possibly differs from the main component (M) in an amide residue. The two slower components (S, and S,) have the same amino acid composition as M, but contain a carbohydrate moiety. That of S, differs from that of S, by one sialic acid residue. These observations, although controversial at the time, were later confirmed by Proctor et al. (1974). The heterogeneity reported for other ruminant a-lactalbumins (Table 11) probably involves, at least in part, glycosylation. The rabbit and rat proteins are unusual in that the main protein is a glycoprotein in each case. The rat protein is unique, furthermore, in having an extension of the peptide chain at the carboxy-terminal end, giving a total of 140 residues versus the usual 123 residues. A necessary, but insufficient, property for a protein to be an alactalbumin is its ability to act as a specifier in the lactose synthase system. There is, at present, controversy as to whether a true a-lactalbumin occurs in the milk of monotremes. Thus, it is essential to have good methods for the determination of galactosyltransferase and lactose synthase activities. They must enable the detection of low levels of activity. This is

LYSOZYME AND a-LACTALBUMIN

191

especially important in considering the possibility of some lysozymes exhibiting weak specifier activity. It is beyond the scope of this article to make a detailed critical evaluation of available methods. Nevertheless, it is important to make several comments. In general, the determination of lactose synthase and galactosyltransferase activities will be necessary on milk samples of the species being studied during the course of fractionation, and on the isolated alactalbumin (or lysozyme). Regardless of the method used, a high-quality galactosyltransferase and a reference a-lactalbumin (usually bovine) will be necessary. It is our experience that some, although not all, commercial preparations of galactosyltransferase and a-lactalbumin are not satisfactory. The latter is frequently impure. It may contain substantial amounts of lactoferrin; indeed, one group described the isolation of lactoferrin from commercial a-lactalbumin as a convenient method of preparation of the former (Castellino et al., 1970). Thus, in general, it is preferable to use high-quality laboratory preparations. Until recently, methods for determination of both activities were essentially of two types: the determination of UDP formation enzymatically by a spectrophotometric method and the determination of the incorporation of UDP[U-14C] galactose into ["C] lactose. The first type is limited to purified systems, since crude systems catalyze the endogenous oxidation of NADH (Brodbeck and Ebner, 1966). The effects of varying conditions of pH and concentrations of substrate (glucose or N-acetyllactosamine), UDP-galactose, and MnC1, on the I4C incorporation determination of lactose synthase and galactosyltransferase in both crude systems and purified proteins have been studied, for example, by Fitzgerald et al. (1970b). The use of UTP in the inhibition of interfering hydrolase activity was discussed by McGuire (1969). In more recent procedures, products (including degradation products) and unused UDP-galactose are separated by high-voltage electrophoresis, and the substrate may be 3H-or 14C-labeled(Ram and Munjal, 1985). Hopper and McKenzie (1974), in their study of the possible ability of echidna lysozyme to act as specifier protein for the production of weak lactose synthase activity, found that the conditions of the determination needed modification for these purposes. A preliminary study was made by H. A. McKenzie and V. Muller (unpublished observations) of optimum conditions for such determinations. However, it was evident that much further work was necessary. The effect of lipids on galactosyltransferase activity has been studied by Mitranic and Moscarello (1980). More recently, Hymes and Mullinax (1984) introduced the use of HPLC in the determination of galactosyltransferase activity. This method

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HUGH A. MCKENZIE AND FREDERICK H.WHITE, JR.

does not require radioactive substrates. Further, it permits the use of saturating levels of UDP-galactose and the monitoring of side reactions. Thus, its further development looks promising for studies of systems exhibiting weak activity.

IV. THREE-DIMENSIONAL STRUCTURE OF LYSOZYME A . X-Ray Crystal Structure of Domestic Hen Egg-White Lysozyme

As indicated in Section I, the determination of the three-dimensional structure of domestic hen egg-white lysozyme was the first elucidation of the X-ray crystal structure of an enzyme (Blake et al., 1962, 1965, 1967a; Phillips, 1966, 1967). One reviewer stated, “up until that time very little was known about its catalytic properties” (Creighton, 1984). Actually, prior to this work a good deal of important information on the nature of the linkages attacked by lysozyme had accumulated due to the important work by Ghuysen, Salton, and others. The X-ray studies indicated the nature of the active site of the enzyme and the mode of binding to inhibitors and substrate. The studies were also important in that they demonstrated the first example in a globular protein of the P-sheet structure and differences from the protein structures previously determined: myoglobin and hemoglobin. The elucidation of the structure was considerably facilitated by the determinations made in two laboratories (those of Canfield and Jolles) of the amino acid sequence, emphasizing the critical importance of adequate sequence information for X-ray studies (the electron density map did not show the individual atoms separately resolved). In summary, the main structural features of the domestic hen eggwhite lysozyme molecule (see Fig. 3) are: 1. The molecule has approximately the shape of a prolate ellipsoid, 4.5 x 3.0 x 3.0 nm (no allowance being made for bound water). It has

a deep cleft on one side. The cleft divides the molecule roughly into two lobes. The first consists of the two ends of the chain (residues 1-39 and 85- 129), while the second (comprising residues 40-84) is rather sheetlike and consists of residues either in the outer surface or lining the cleft. 2. In contrast to myoglobin and hemoglobin, lysozyme has a fairly small proportion of helix and reasonably long stretches of chain with essentially irregular conformation. Several parts of the chain have, as already mentioned, an extended conformation closely similar to the p sheet seen in fibrous proteins.

LYSOZYME AND a-LACTALBUMIN

193

FIG.3. Perspective drawing of the main-chain conformation of domestic hen egg-white lysozyme. The view is an elevation from the active-site side of the molecule (Imoto et al., 1972). Only the positions of the a-carbon atoms are shown. An instructive colored painting by I. Geis of the original three-dimensional model of lysozyme is reproduced in the early review by Phillips (1966). Of historic interest is the drawing of the model by the late Sir Lawrence Bragg (reproduced by Blake et al., 1965, and Phillips et al., 1987). It is to be noted that Bragg’s diagram is a free-hand drawing and not an accurate computergenerated representation of the molecule. (Reproduced with permission from Imoto et al., 1972.)

3. In detail: The first lobe (residues 1-39 and 85- 129)contains four helices that are close to the Pauling-Corey a-helix type, and one singleturn 310-typehelix. There are short stretches (each five to nine residues) of backbone loops and turns connecting the helices. Three a helices (helix A, residues 4- 15; helix C, residues 88-99; helix D, residues 108-115) are on the protein surface and are partially exposed to solvent. The a helix (B) consisting of residues 24-36 is totally buried. The 310helix (residues 119- 124) is partially exposed to solvent. The second lobe (residues 40-84) contains a three-stranded antiparallel P-pleated

194

H U G H A. MCKENZIE AND FREDERICK H. WHITE, JR.

sheet (residues 42-60), a small p sheet (residues 1-2 and 39-40), and a single-turn 3,, helix (residues 79-84). There is a long coiled-loop region, residues 61-78, between the large p sheet and the 3,, helix. Residues that line the cleft include p-sheet residues 43, 44, 46, 52, and 56-59, helix B residue 35, the loop connecting helices C and D (residues 98 and 101- 103), helix D residues 107- 110 and 112, and residues 62, 63, and 73. 4. Cystine bridges occur between residues 6 and 127, 30 and 115, 64 and 80, and 76 and 94. The first two pairs have negative torsion angles, but the last two pairs have positive angles. All are-in the range 100” ? 10”. 5. Following the original work by Kauzmann on hydrophobic interactions and the determinations of the structures of myoglobin and hemoglobin, it was stated, and is still stated frequently (despite evidence to the contrary), that hydrophobic residues are buried in the interior of proteins and hydrophilic residues are exposed to solvent water. It was first shown by Klotz (1970; see also Lee and Richards, 1971) that a substantial proportion of the exposed solvent-accessiblesurface area of proteins is composed of nonpolar groups. This matter has been stressed in lectures for many years by one of the authors (H. McK.) (for a discussion of various approaches to this problem, see Edsall and McKenzie, 1983). In the case of lysozyme, a substantial proportion of the hydrophobic residues Leu, Val, Ile, Ala, Gly, Phe, Tyr, Trp, Met, and Pro are either fully exposed to solvent or at least have some atoms that are solvent accessible. Examples of “hydrophobic” residues that are “surface” exposed are Val-2, Phe-3, Leu-17, Phe-34, Leu-75, Trp-123, Pro-’70, and Pro-79, with Trp-62, Trp-63, Ile-98, Trp-108, and Val-109 being on the surface of the cleft. Examples of the least-exposed ionizable side chains are Asp-66, Asp-52, Tyr-53, His-15, and Glu-35. The above summary is based on the structure of the tetragonal crystalline form of domestic hen egg white lysozyme determined at 0.2-nm (2 A) resolution. A refined high-resolution (0.15 nm, 1.5 A) study has been made by Handoll et al. (unpublished observations, quoted by Post et al., 1986). This study includes refinement of the positions for interior and surface water molecules. As shown in Table 111, other crystalline forms have been isolated and studied by X-ray crystallography. Joynson et al. (1970) studied the triclinic and tetragonal forms of hen egg-white lysozyme; Moult et al. (1976) studied the triclinic form; Hogle et al. (1981) compared monoclinic, triclinic, and tetragonal forms; and Artymiuk et al. (1982) studied the monoclinic and orthorhombic forms (see also Table 111). The results from these studies have shown essen-

LYSOZYME AND LY-LACTALBUMIN

195

tially the same conformational structure for all of these crystalline forms. However, it is important to realize that the lysozyme molecules are more closely packed in the triclinic crystals than in the tetragonal crystals. This may account for the fact that the apparent thermal factor ( B ) ' is lower in the triclinic form (B = 8) than in the tetragonal form (B = 15). In several instances long flexible side chains have very different conformations in the two structures (e.g., Arg-14, Lys-33, Phe-38, Arg-61, Arg-73, Arg-114, and Arg-128). There are also some differences in main-chain conformation, especially in the p-loop region between residues 44 and 50. Jolles and Berthou (1972) observed that tetragonal crystals of lysozyme were unstable above 25"C, especially at physiological temperatures, and transformed into orthorhombic crystals which are stable u p to 55°C (see also Berthou and Jollts, 1974). Berthou et al. (1983) found that, although the conformations obtained from orthorhombic and tetragonal forms are similar, there are differences caused by crystal contact. Thus, Trp-63 and Pro-71 are much better ordered than in the tetragonal form, where they are exposed to solvent. These differences may account for the observed difficulty of inhibitor binding in the hightemperature crystalline form, but do not seem to reflect the behavior of lysozyme in solution at the same temperature. B . Mechanism of Cell Lytic Action

We have already seen that lysozyme is a glycosidase hydrolyzing the glycosidic bond between C-1 of N-acetylmuramic acid (NAM) and C-4 of N-acetylglucosamine (NAG) of bacterial cell wall polysaccharide (Section 11,A).The polysaccharide chitin, found in crustacean shells, consists only of NAG residues joined by p( 1+4)-glycosidic links. It is also a substrate for lysozyme. The identification of the substrate binding site and the mechanism of the catalysis were not immediately evident from the original X-ray studies of lysozyme. One approach would be to apply the difference Fourier method (see Blundell and Johnson, 1976) to elucidate the structure of the enzyme-substrate complex during catalysis. Such an approach is impractical at room temperature because of the slow rate of X-Ray results provide important information regarding molecular and atomic motions, through determination of the thermal factor ( B ) ,which gives a measure of the mean square (harmonic) displacement (3) of an atom or group from its equilibrium position. The two are related by the Debye-Waller equation: B = 8+$. A highly mobile protein side chain may have a B value as high as 40 A2,corresponding to a mean square displacement of 0.5 A2 (see also the discussion in Section IV,D).

TABLE 111 Crystallografhic Data for Lysozyme and a-Lactalbumin Cell dimensions

Source Lysozyme Domestic hen egg white

Crystal form

Tetragonal

Orthorhombic Monoclinic

Growth conditions (medium, pH, temperature)

-0.9 M NaCI, pH 4.5-4.7, 18°C 0.3-1.5 M NaClb, pH 4.3, 18°C 0.5-1.1 M KClb, pH 4.3, 18°C 0.5-1.1 M NH4Clb, pH 4.3, 18°C 0.4-1.1 M MgClz', pH 4.1, 18°C 0.5-1.2 M ammonium citrateb, pH 4.7, 18°C 0.9-1.5 M NH40Acb, pH 4.5, 18°C 1.1-1.2 M NaHzP04*, pH 4.5, 18°C 0.9 M NaCI, pH 10, RTc 0.36 M N a N Q , H N Q , pH 4.5, RT 0.77 M Na2SO4, 0.5 M NaOAc + HzS04, pH 4.5, RT 0.2 M Nal, pH 4.5, RT 0.075-0.20 M KSCNb, pH 4.5,

Space group

P43212

P212p21 p2 I

18°C

Triclinic Tortoise (Tnmyx gangetuus) Cuvier egg white Human urine (leukemic)

Moll asym. unit Refs."

a (")

(i) (A) (i)

p

y

(")

(")

79.1 79.2 79.2 79.2 79.2

79.1 79.2 79.2 79.2 79.2

37.9 38.0 38.0 38.1 37.9

90 90 90 90 90

90 90 90 90 90

90 90 90 90 90

1 1 1 1

78.8 79.2 79.0 56.3 27.9

78.8 79.2 79.0 65.2 63.1

38.3 37.9 38.1 30.6 60.6

90 90 90 90 90

90 90 90 90 90.5

90 90 90 90 90

1 1 1 1 2

28.6 28.1

63.0 63.1

61.6 60.4

90 90

93.5 91.0

90 90

2 2

28.1

63.0

60.4

90

90.4

90

2

4

108.8 90

111.5 90

1 1

6

90

90

1

7,s

b

Orthorhombic

0.24 M N a N Q , 0.025 M NaOAc, pH 4.5, RT NaZHFQ4, KHzP04, pH 6.6, RT

P1 P212121

27.5 58.0

32.0 58.9

34.4 43.1

88.5 90

Orthorhombic

7 M NH4N03, pH 4.5, RT

P212121

57.1

61.0

32.9

90

1-3 4

1

2 3

3,5

a-Lactalbumin Baboon (Papio qynocephaluc) milk Human milk

Goat (Capra hircw) milk

Orthorhombic (spheroidal) Orthorhombic Orthorhombic

Monoclinic Cow milk

Triclinic Tetragonal Monoclinic Trigonal I Hexagonal Trigonal I1

-1 u satd. (NH4)2SO4 + 1 u 0.2 M PO;, pH 6.8, RT 1.8 M (NH4)2S04, 01. M PIPES, -0.006 M CaCb, pH 6.5, 35°C -1 u satd. (NH4)2SO4 + 1 u 0.2 M PO4 (or 0.1 M Tris-HCI), pH 6.6, RT, “high salt crystals” Water + minimum satd. NaCI, p H 5.3, RT, “low salt crystals” 0.5 satd. (NH4)2S04. 0.1 M Pod, p H 6.6, 25°C 0.5 satd. (NH4)2S04. 0.1 M PO4, p H 6.6, 4°C 0.5 satd. (NH&S04, 0.2 M PO,, p H 6.6, 4°C 1.9 M (NH4)2S04, 0.2 M PO,, p H 6.5, 4°C 1.8 M (NH&S04. 0.2 M PO,, p H 6.5, 35°C 1.9 M (NH4)2S04, 0.1 M PIPES, 0.01 M CaCI2, pH 6.5, 35°C

33.6 35.5* 33.6

69.6 69.1d 69.9

47.0 46.1d 47.3

90 90 90

90 90 90

90 90 90

1 1 1

67.6

109.7

68.9

90

90

90

2

45.0

89.0

32.1

90

92.6

90

2

94.7

122.9

117.9

90

116

91

32

119.6

119.6

153.2

90

90

90

8

140.7

196.7

63.2

90

111

90

24

57.4

57.4

75.0

90

90

90

I

94.0

94.0

67. I

90

90

90

1

93.7

93.7

66.9

90

90

90

2

“References: (1) Alderton and Fevold (1946),(2) Palmer et al. (1948), (3) Steinrauf (1959), (4) Ries-Kautt and Ducruix (1989). (5) Moult et al. (1976), (6)Aschaffenburg al. (1980). (7) Osserman and Lawlor (1966), (8) Banyard et al. (1974), (9) Aschaffenburg et al. (1979), (10) Fenna (1982b), (11) Aschaffenburg el al. (1972h), (12) Aschaffenburg et al. (1972a), (13) Fenna (1982a). b0.005 M in NaOAc.
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HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

diffusion of substrate into the enzyme crystal compared to the rate of conversion of bound substrate into product. Another approach is to cool the crystal (e.g., - 20 to - 50°C) to slow the catalytic process and to examine the enzyme-substrate complex under the low-temperature conditions. The feasibility of this cryoenzymological approach has been discussed by Douzou (1977). As far as we are aware, the only attempt to determine the feasibility of this method for lysozyme is that by Douzou et al. (1974). Blake et al. (1967b; see also Johnson and Phillips, 1965) used another approach: a difference electron density study of the complex of lysozyme with an “unreactive” analog of the substrate, compared with lysozyme. Oligomers of N-acetylglucosamine consisting of fewer than five residues either are not hydrolyzed or are hydrolyzed only very slowly, but they do bind to the enzyme. Tri-N-acetylglucosamine (tri-NAG) is a potent competitive inhibitor of lysozyme. Thus, the &-ray study (2 resolution) of the tri-NAG-lysozyme complex revealed the active site and the interactions involved. The binding of the tri-NAG was shown unequivocally to be in the enzyme cleft. The three residues (at positions designated A, B, and C) filled only one-half of the cleft. There was room for three additional residues (at positions D, E, and F); this is an important point because hexa-NAG is rapidly hydrolyzed by lysozyme. The positions (D, E, and F) of the three additional rings were deduced from model building. This is shown in Fig. 4, and a schematic diagram is given in Fig. 5. Important evidence from chemical studies considerably assisted the crystallographic work. Rupley (1967) showed that the relative rates of hydrolysis of n-mers of NAG (trimer taken as unity at M substrate concentration) are 1 (trimer), 8 (tetramer), 4 x 103 (pentamer), 3 x 104 (hexamer), 3 x lo4(octamer); that is, there is a dramatic increase in rate from (NAG)4to (NAG)5,with a further increase for (NAG)6,but a negligible change thereafter. Since (NAG)s is stable, the A-B and B-C bonds cannot be the site of cleavage for (NAG)6.Furthermore, Rupley found that lysozyme cleaves the hexamer into a tetramer and a dimer, the latter two residues being at the reducing end of the oligomer; that is, the cleavage is between residues D and E (see also Shrake and Rupley, 1980). If we consider the cleavage of the cell p( 1+-4)-glycosidiclinkage, it will be recalled that this involves the linkage between NAM and NAG residues and not that between NAG and NAM. In their model-building experiments Blake, Johnson, Phillips, and co-workers found that residue C could not be NAM because there was no room for the lactyl side chain. T h e same was true for residue E, so that only residues B, D, and F could be NAM. Since cleavage of the linkage between residues B and C has

FIG. 4. The active site of hen egg-white lysozyme, showing the mode of binding of a hexasaccharide (hexa-N-acetylchitohexaose),as proposed and developed by Blake, Johnson, Phillips, and co-workers. The sugars in sites A-C are based on the crystallographic determination of the binding of tri-N-acetylchitotriose, and those in sites D-F are based on model building. Cleavage of the hexasaccharide is considered to take place between D and E (see text). The monosaccharide N-acetylglucosamine was found to bind to site C in two slightly different orientations, one of which corresponds to the anomer (as here), the other corresponding to the binding of the a anomer (see Fig. 1B of Perkins et al., 1981). (Reproduced with permission from Perkins et al., 1981.)

200

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

Trp-62 Trp-63

Asn-103

Asn-59 Ala-107 Asn-46 Tr p-108 Val-1 09

0

Asn-37

0

Phe-34

Arg-114

FIG. 5. The binding sites for the hexasaccharide to lysozyme, indicating residues implicated in the catalytic cleavage.

already been excluded, it was evident that the linkage between residues D and E was highly likely to be the one cleaved, and this is supported from the above chemical studies on the cleavage of (NAG)G. Further information was obtained from chemical studies in which the enzyme-catalyzed hydrolysis was performed in the presence of water enriched with l 8 0 . It was concluded that the bond split is that between C-1 of residue D and the oxygen of the glycosidic linkage to residue E. It is evident that any detailed mechanism of the catalytic action of lysozyme in the hydrolysis of glycosides would involve the role for proton donors and/or acceptors. The most plausible residues near the cleavage site are Asp-52 and Glu-35. The Asp residue lies in a polar environment, where it is a hydrogen bond acceptor in a complex network of hydrogen bonds. Because of its location, it has virtually a normal pK, of 3.5 ? 0.2. At pH 5.0, which is near the optimum pH value of hydrolysis of chitin by lysozyme, Asp-52 is in the ionized form. The Glu residue lies in a nonpolar region, has an increased pK, of 6.3 k 0.2, and would be largely un-ionized at pH 5.0. The mechanism involving these residues, as developed by Blake,Johnson, Phillips, and co-workers over the period 1965- 1972 (see also Vernon, 1967)is as follows: A hydrogen ion is donated from the carboxyl of Glu-35 to the glycosidic oxygen between the rings in sites D and E, resulting in the cleavage of the bond between C-1 of the ring at D and the glycosidic oxygen, creating a positive charge on C-1. This transient species

20 1

LYSOZYME AND OI-LACTALBUMIN

is designated as an oxocarbonium ion, which may be stabilized by the negative charge of Asp-52. With the removal of the NAG dimer from sites E and F, the carbonium ion intermediate reacts with an OH- of surrounding water. Also, Glu-35 becomes protonated, tetra-NAG moves from sites A, B, C, and D, and another round of catalysis may then occur. An essential feature of this proposed mechanism (Fig. 6, Scheme 11) is the distortion of the NAG residue at site D from its normal chain conformation. The resultant twist boat conformation enables stereoelectronic assistance to be obtained from ring oxygen 0, in the transition state, leading to cleavage of the exocyclic C,-0, bond. The overall mechanism has received support from a good deal of experimental work (see, e.g., Imoto et al., 1972; Ford et al., 1974; Banerjee et al., 1975; Rosenberg and Kirsch, 1981). When tetra-NAG is treated with lysozyme, some hexa-NAG and diNAG are formed. This reaction, transglycosylation, was first proposed by Maksimov et al. (1965) to account for hydrolysis of tri- and tetrasaccharides, during which an insoluble chitinlike carbohydrate was formed. Thus, short-chain oligosaccharides are cleaved by lysozyme, with the products transferred to longer-chain oligosaccharides, which are subsequently broken down. Much work has focused on this phenomenon,

0 Glu-35

Scheme I

\

H

\

Glu-35

Scheme 11

0

5\

Glu-35 -0

FIG. 6. Proposed reaction mechanisms by which lysozyme catalyzes the cleavage of a polysaccharide substrate. Scheme I is that proposed by Post and Karplus (1986) and indicates the possibility of cleavage with no prior assumption of distortion of the sugar ring at site D. Scheme I1 is that originally developed by Blake, Johnson, Phillips, and co-workers and involves ring distortion as a critical step. (Reproduced with permission from Post and Karplus, 1986.)

202

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

which is well treated in the review by Imoto et al. (1972). While transglycosylation complicates kinetic studies of lysozyme catalysis, it has also indicated general support for the above mechanism. However, a good deal of recent experimental and theoretical work does not lend support for the distortion of the sugar ring at site D. In an X-ray study [0.25-nm (2.5 hi) resolution], Kelly et al. (1979) found no direct evidence for distortion of ring D in the complex between lysozyme and the trisaccharide NAM-NAG-NAM, bound in subsites B, C, and D. They concluded that it was more likely that they were looking at a Michaelis complex. In a series of papers, Pincus, Scheraga, and co-workers (see, e.g., Pincus and Scheraga, 1981) studied the conformational energies of complexes of lysozyme with oligomers of NAG and copolymers of NAG and NAM. T h e search of the conformational space at the active site of lysozyme and the minimization of the conformational energies of the complexes indicated: the hexasaccharide (NAG), binds preferentially on the “left” side of the lysozyme cleft. The fourth residue from the nonreducing end is bound in the chair form at a D site close to the surface of the cleft, somewhat removed from Glu-35 and Asp-52. The binding of the fifth and sixth residues of (NAG), at sites E and F involved residues such as Arg-45, Asn-46, and Thr-47. This D-site binding mode is in accord with the solution studies by Schindler et al. (1977) and the X-ray studies by Kelly et al. (1979). There were two other structures, but of somewhat higher energy. One of them had a distorted conformation, but the best contacts, at sites E and F, were on the “right” side of the cleft. There were contacts at site F with residues Asn-113 and Arg-114, the structure being similar to the model discussed by Imoto et al. (1972). T h e hexasaccharide (NAG-NAM),-(NAG), was also found to bind on the “left” side of the cleft. In contrast, the alternating copolymer (NAG-NAM)3 was bound with its F-site residue on the “right” side, residues such as Phe-34 and Arg-114 being involved (the lacteal side chain of NAM prevents F-site binding on the “left” side). The calculations indicated that the highest affinity of the disaccharide NAG-NAM is for sites C and D and the “right”-side sites E and F, in agreement with the experimental study by Sarma and Bott (1977). T h e conformational energy calculations received support subsequently from two types of experimental study by Smith-Gill et al. (1984). T h e affinity of ring-necked pheasant lysozyme, in which Asn and Arg at positions 113 and 114 are replaced by Lys and His, respectively, is the same for (NAG), as that of domestic hen egg-white lysozyme (i.e., the right side is not involved). They also showed that a monoclonal antibody bind-

LYSOZYME AND (Y-LACTALBUMIN

203

ing specifically to an epitope including residues Arg-45, Asn-46, Thr-47, Asp-48, and Arg-68 on the “left” side of hen egg-white lysozyme, is competitively displaced by (NAG), and (NAG),, but not by NAG, (NAG),, o r (NAG), [i.e., the terminal residues of (NAG)s and (NAG), bind to the “left” side]. In a molecular dynamics study of native and substrate-bound hen eggwhite lysozyme, Post et al. (1986) found the structural features analyzed agreed well with the results of X-ray studies at 0.15-nm ( I .5 h;) resolution, except for some surface residues. Appreciable differences were found in residue mobilities between the simulations of the native and substrate-bound states in the region of the enzyme that is in direct contact with the substrate and in a region that is distant from the active site cleft. This study enabled Post and Karplus (1986) to develop a case for an undistorted ring at site D in their proposal of an alternative pathway for lysis of the oxygen bridge between rings at sites D and E. An essential feature of their mechanism is that no twist-boat (“half-chair”) conformation for ring D is necessary, assuming certain minor rearrangements among residue side chains are made in the structure of lysozyme. Another essential feature is that, with this undistorted substrate in place within the binding site cleft, the endocyclic bond between the oxygen and C, of the ring at site D (Fig. 6, Scheme I), rather than the exocyclic bond originally proposed by Blake et al. (1967b), breaks first. T h e initial step in the reaction is protonation of ring 0, by Glu-35, followed by cleavage of the endocyclic bond C- 1-0-5 with formation of the oxocarbonium ion intermediate, stabilized by Asp-52. Hydrolysis, cleavage of the C,-OS bond, and ring closure give rise to the reaction products. T h e importance of entropic, rather than enthalpic, contributions, as in the “classic”mechanism, has been discussed by Post and Karplus (1986), who carefully stress that, although their mechanism is in accord with experimental results, it is only suggestive at this stage. Genetic engineering of hen egg-white lysozyme has been used by Kirsch et al. (1989) as an approach to studying the structure-function relationships of lysozyme. Thus, they offer evidence from site-directed mutagenesis of cloned lysozyme (expressed in yeast), that Asp-52 and Glu-35 are vital for the expression of lysozyme. However, it is curious that conversion of Asp-52 to the amide resulted in a form of the enzyme that still had 5% of the normal activity. Conversion of Glu-35 to the amide, on the other hand, resulted in a lysozyme that was devoid of all activity. It was demonstrated by mutagenesis of Asp-101 to Gly that the ionization of this residue contributes thermodynamically to the association of lysozyme with the inhibitor chitotriose.

204

HUGH A. MCKENZIE AND FREDERICK

H. WHITE, JR.

Dao-pin et al. (1989) stressed that the enzymatically catalyzed hydrolysis of polysaccharides proceeds at more than five orders of magnitude faster than that for model compounds mimicking the substrate in the active site of the lysozyme. Although many workers have stressed that electrostatic interactions of specific residues with the substrate are an important feature of the mechanism, Dao-pin et al. suggest, rather, on the basis of results obtained by classical electrodynamics, that the charge distribution of the enzyme as a whole is the important feature. C . Structures of Other Lysozymes

In addition to the studies of various crystalline forms of domestic hen egg-white lysozyme, the structures of human and tortoise egg-white lysozymes have been determined (for crystal data see Table 111). Artymiuk and Blake (1981) refined the structure of the human enzyme to 1.5 8, resolution. The main objectives of this study were to determine the extent of differences in structure from that of the hen egg-white protein, to discover the location of water molecules, and to test the validity of the method of restrained refinement. The particular restrained leastsquares approach to refinement described in their paper appears to have been validated. The two proteins were found to be closely homologous, but there were small differences (e.g., in a helices), details of which can be obtained from consulting their paper. The X-ray refinement of the tortoise enzyme has been used by Blake et al. (1983) primarily to study the location of water molecules, as discussed in Section IV,D. The egg white of the Australian black swan, Cypus atratus, contains two forms of lysozyme c and g types. The X-ray structure of the g type has been determined by Isaacs et al. (1985)at 0.28-nm (2.8 A) resolution. A comparison of the structures of chicken egg white, goose egg white (g type), and bacteriophage T4 types of lysozyme has been made by Weaver et al. (1985), and the evolutionary relationships have been discussed in light of this study. This work is discussed in Section X.

D . Water in Lysozyme Crystals The total volume of protein crystals is 25-65% water, and problems that are not yet completely resolved are the nature and location of the water molecules. However, much light is being shed on the problem by high-resolution (0.12-0.18 nm; 1.2- 1.8 A) X-ray diffraction studies, supplemented more recently by neutron diffraction studies (Kossiakoff, 1985). Edsall and McKenzie (1983) have made a tentative classification

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of the major categories of water associated with proteins, summarized as follows: 1. Outside the immediate neighborhood of the protein surface, the

water is essentially bulk water (modified only by the presence of salt ions and small organic molecules). 2. Water at the protein surface consists of three subcategories: (a) highly mobile water molecules adjacent to nonpolar surface atoms, (b) somewhat less mobile water molecules hydrogen bonded to polar groups of Ser and Thr, C=O and NH groups of the peptide chain, and (c) water molecules that are probably more retarded around groups carrying a formal charge (Lys, Arg, Asp, Glu, etc.). 3. Internal water molecules within the folded peptide chain. 4. Water molecules at the interface of subunits, for those proteins consisting of subunits. T h e results of the study by Blake et al. (1983) of human and tortoise egg-white lysozymes at high resolution is broadly consistent with the above classification. (In all such studies certain assumptions are made regarding the B factors and occupancy and displacement.) Blake et al. found that, although the tortoise lysozyme crystals had -650 water molecules per protein molecule versus 350 for the human protein molecule, the numbers of ordered water molecules were similar for both (i.e., 128 versus 140) (cf. also triclinic and tetragonal forms of domestic hen egg-white lysozyme; 110 and 140, respectively). Their results are summarized in Table IV. On average, protein oxygen atoms interact with twice as many water molecules as protein nitrogen. In conformity with results with other proteins, twice as many water molecules are hydrogenbonded to peptide C=O groups as to peptide NH. It is evident from the results of Blake et al. that, as a group, the charged side chains-Asp, Glu, Lys, and Arg-have the highest solvation. (Other results are given in detail in the original paper.) Neutron diffraction crystallographic studies of the dynamics and hydration of lysozyme are discussed in Section XI. In an investigation of the role of water in enzymic catalysis, Brooks and Karplus (1989) chose lysozyme for their study. Stochastic boundary molecular dynamics methodology was applied, with which it was possible to focus on a small part of the overall system (i.e., the active site, substrate, and surrounding solvent). It was shown that both structure and dynamics are affected by solvent. These effects are mediated through solvation of polar residues, as well as stabilization of like-charged ion pairs. Conversely, the effects of the protein on solvent dynamics and

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HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

TABLE IV Protein- Water Interactions in Human and Tortoise Lysozymes a Property No. of ordered water moleculesb No. making hydrogen bonds to bound water6 No. making at least two hydrogen bonds with protein* (b) No. making one hydrogen bond with protein* (c) Total no. bound to protein* [(b) + (c)] % Bound to peptide CO % Bound to peptide NH % Bound to side chains % Bound to protein oxygens % Bound to protein nitrogens Mean H20-0 distance (A) Mean H2O-N distance (8) Mean H 2 0 - H 2 0 distance (8)

Human

Tortoise

140 35 35 70 105 42 18 40 68 32 2.83 +- 0.15 2.96 t 0.12 2.82 2.0.23

128 19 33 76 109 44 18 38 64 36 2.81 2 0.15 2.96 t 0.17 2.84 t 0.23

“From work by Blake el al. (1983). Per molecule of protein.

structure were also observed to be significant. In particular, the water surrounding apolar groups is less mobile than bulk water, or the water solvating polar groups.

V. THREE-DIMENSIONAL STRUCTURE OF a-LACTALBUMIN

A. Mo&h for the Three-Dimnsional Structure of a-Lactalbumin (Based on Sequence Homology with Lysozyme) Because of the high level of identity in amino acid sequence between lysozyme and a-lactalbumin (see Fig. lo), it was inevitable that interest turned to the three-dimensional structure of a-lactalbumin when the structure of lysozyme was determined in 1965 by the group at the Royal Institution. However, there were unforeseen difficulties in the direct experimental determination, as discussed below. Hence, attention was directed to models for the a-lactalbumin structure based on the coordinates for lysozyme and on energy minimization programs. I . Model of B r o w et al. ( I 969) A wire skeletal model of lysozyme was constructed and then modified to accommodate the a-lactalbumin sequence, by changing side chains that differ between them and by rearranging the main chain to accommodate the various deletions.

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Browne et al. (1969) concluded that the differences between the two sequences were compatible with their having similar conformations; that is, hydrophobic side chains are usually replaced by other hydrophobic side chains, and a change in one residue is often accompanied by a compensating change in a neighboring residue. All of these changes were so readily accommodated, without producing major rearrangement of the main-chain conformation, that the proposed model for a-lactalbumin seemed likely to be substantially correct. The effect of the changes in the upper part of the substrate cleft appears to be that sites A and B are blocked off, largely a consequence of the replacement of Ala-107 (see Section VI) in lysozyme with Tyr (or His in the rabbit). These changes make it unlikely that sites A and B would remain attractive saccharide binding sites in a-lactalbumin. However, as discussed in Section VI, 107 is not always Ala in lysozyme. The lower part of the cleft, where residues E and F bind in lysozyme, is changed both in topology and in the nature of surface groups. While it appears from this model that this part of the cleft could bind saccharides, the precise mode of binding would be expected to differ from that in lysozyme. Residue 52, Asp, is invariant in lysozyme, the equivalent residue in a-lactalbumin being Glu-49. Residue 35 is Glu in lysozyme, but the equivalent residue in a-lactalbumin is variable. However, if residues 32-36 are rearranged to give maximum sequence identity: Hen egg-white lysozyme

32 33 34 35 35a 36

Ala LYS Phe Glu

J. Ser

Bovine a-lactalbumin Thr

30

I Phe His Thr Ser

31 32 33 34

then the equivalent bovine a-lactalbumin residue for hen egg-white lysozyme residue 35 (Glu) is His-32. This residue could assume the function of Glu-35. On the basis of this model, it might be anticipated that a-lactalbumin and lysozyme have similar biological functions, as well as similar conformations. 2. Calculations of Lewis and Scheraga (1971) The probability that particular residues start or end in helix was calculated for bovine a-lactalbumin and hen egg-white lysozyme. There is a one-to-one correspondence of location of helical regions, predicted and found for lysozyme and predicted for a-lactalbumin. Thus, the

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WHITE, JR.

overall impression is one of great similarity between the two proteins. The helical content in the carboxy-terminal region, however, is predicted to be greater for a-lactalbumin.

3. Model of W a r n et al. (1974) When Warme et al. (1974) used their method for minimizing the total potential energy of a protein, in the examination of hen egg-white lysozyme, they found a low conformational energy that still maintained a close resemblance to the original X-ray structure. Then they adapted their procedures to compute a low-energy conformation for a-lactalbumin, based on the known structure of lysozyme. The following points especially stand out. In the cleft region the His-32 side chain points down and away from the cleft region and does not occupy the position corresponding to Glu35 of lysozyme. This differs from the conclusion of Browne et al. (1969), who, as we have seen, suggested that His-32 might substitute for Glu-35 in the active site. In the Warme et al. (1974) model residue 33, T h r in bovine a-lactalbumin, but not invariant in other species, appears to correspond more closely in position to Glu-35 in lysozyme. In the upper part of the cleft region, Tyr-103 tends to block Trp-60 and Trp-104 from contact with substrates. This agrees with the finding by Browne et al. (1969), that is, that sugar binding sites A and B of lysozyme are blocked in a-lactalbumin. Reactivities of Tyr residues were found to be somewhat difficult to interpret in terms of past experimental observations, although the model and the experimental results were in “reasonable” agreement. Warme et al. (1974) concluded, for example, that Tyr-18 and Tyr-103 would be those most easily acetylated with acetylimidazole (Kronman et al., 1972a), while Tyr-36 and Tyr-50 would react more slowly. Castellino and Hill (1970) reported that the Met is readily accessible to reagents, in agreement with the model. Also in agreement were the reactivities of His residues. Thus, with iodoacetate, carboxymethylation proceeds in the order His-68 > His-32 > His-107. Helical contents are much the same as those reported for lysozyme (Robbins and Holmes, 1970; Bare1 et al., 1972). Carboxyls are, in general, more exposed in a-lactalbumin than in lysozyme (Lin, 1970), in agreement with the model. Immunochemical differences observed experimentally (i.e., no crossreactivity) are not incompatible with the model, which shows many surface differences with lysozyme. The greatest difficulty with this model, as with that of Browne et al. (1969), lay in predicting the structure of the carboxy-terminal end of

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a-lactalbumin: neither of the two groups was able to suggest a unique structure for this part of the molecule. Indeed, the elucidation of this structure had to await the solution of the X-ray structure.

B . X-Ray Crystal Structure of Baboon Milk a-Lactalbumin Attempts to produce crystals of bovine a-lactalbumin from the milk of Western dairy breeds of cattle for X-ray crystallographic studies met with appreciable difficulties. While crystallization from concentrated ammonium sulfate was not difficult, the crystals were very small. Thus, Aschaffenburg et'al. (1972a) were led to the study of crystallization of the goat milk protein. Freeze-dried caprine a-lactalbumin was dispersed in water and dissolved by the addition of a minimum volume of saturated NaCl solution, giving a final protein concentration of 10 g dl-I. T h e pH was adjusted to 5.3 and the solution was dialyzed against water at 4°C for several days. The mixture was then warmed to -17"C, resulting in the formation of lozenge-shaped crystals. Attempts to produce heavy metal derivatives of these crystals were not satisfactory. Accordingly, Aschaffenburg et al. (1972b) turned their attention to crystallization from concentrated ammonium sulfate solution. This resulted in crystals that gave a complex diffraction pattern, the additional reflections of which could be eliminated by soaking the crystals in 0.001 M K,PtCl,. Although the caprine crystals looked promising, they proved difficult to analyze. Hence, attention was then directed toward a-lactalbumins of other species, especially the baboon (Papio cynocephalus). Aschaffenburg et al. (1979) found the crystals to be relatively easy to prepare and suitable for structural analysis at high resolution. The turning point in the structural studies came after the work by Hiraoka et al. (1980) revealed that a-lactalbumin is a metalloprotein in which calcium is strongly bound (see also Sections VI and VII). Soon afterward three new crystal forms of bovine a-lactalbumin were isolated by Fenna (1982a), particularly trigonal Form 11. Fenna (1982b) also isolated calcium-containing crystals of human a-lactalbumin suitable for X-ray structural analyses. T h e various crystalline forms of a-lactalbumin are summarized in Table 111. In any event it was the analysis of baboon a-lactalbumin crystals for which the first X-ray crystal structure was produced, initially at 0.6 nm (6 A) and 0.45 nm (4.5 A) (Phillips et al., 1987; Smith et al., 1987). More recently, the structure has been refined at 0.17-nm (1.7-A) resolution, enabling comparison with the high-resolution c-type lysozyme structure (Acharya et al., 1989) (see Fig. 7). As already indicated, difficulties were experienced in the preparation

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HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

C

FIG. 7. The tertiary structure of (a) baboon a-lactalbumin and (b) domestic hen eggwhite lysozyme. (Reproduced with permission from Acharya et al., 1989; based on a program of J. P. Priestle.)

of heavy-atom derivatives of a-lactalbumin. In particular, the attempts to prepare chloroplatinite and bromoplatinite derivatives by the crystalsoaking method were disappointing. Recourse was had to the insertion of a mercury atom in the disulfide bridge linking residues 6 and 120. This involved reduction of cystine residues in the a-lactalbumin crystals soaked in another liquor containing dithiothreitol, and then in another liquid containing mercury(I1) acetate. Although the resultant electron density maps (Smith et al., 1987) did not enable a high resolution of structure, the overall features of the models of Browne et al. (1969) and Warme et al. (1974) were confirmed. Also, the identification of helix B (residues 24-36 in hen egg white lysozyme) in a-lactalbumin enabled resolution of a problem found in the

LYSOZYME AND a-LACTALBUMIN

21 1

earlier work. Although maximum identity could be achieved by deletion at residue 33 (as indicated above) with consequent loss of helix, it had been decided to take a conservative approach and retain the helix. It now appeared that the latter decision was correct. Subsequently, the baboon a-lactalbumin structure was refined at 1.7-A resolution by Acharya et al. (1989). Using the structure of domestic hen egg white lysozyme as the starting model, preliminary refinement was made using heavily constrained least-squares minimization in reciprocal space. Further refinement was made using stereochemical restraints at 1.7-A resolution to a conventional crystallographic residual of 0.22 for 1141 protein atoms. Some features of the refined structure are: 1. The human a-lactalbumin amino acid sequence was used in the refinement since the baboon sequence has not been determined, although it was known from the unpublished work by R. Greenberg to be close to the human sequence. However, it became evident in the course of the X-ray work that there were eight sequence changes in baboon a-lactalbumins (see Section VII,B). 2. The disulfide bridges are similar to those of lysozymes, with the exception of one bridge in echidna lysozymes I and 11, discussed in Section VII,B. 3. There are similarities in the helices and /3 sheets between baboon a-lactalbumin and hen egg-white lysozyme, as summarized in Table V. However, there are important differences, for example, in hen egg-white lysozyme residues 41-60 form an irregular antiparallel @pleated sheet; in this protein a residue is deleted at position 48 (human lysozyme numbering), but two residues are deleted in a-lactalbumin at positions 47 and 48 (human lysozyme numbering). Residue 47 is the most exposed to solvent in the hen egg-white lysozyme and forms part of the irregular p turn. These residues occur in a P-pleated sheet and the deletions are accommodated with minimal disruption to the pleated sheet (see the comparison in Acharya et al., 1989). 4. There are differences in the carboxy-terminal region of a-lactalbumin from lysozyme (see Acharya et al., 1989). This work resolves the inconclusive nature of the earlier models that could not resolve the structure of a-lactalbumin in this region. Also, changes occur in the loop region. 5. Of the 150 water molecules in the a-lactalbumin structure, four have been shown to be internal. Of the two cavities in a-lactalbumin, one small cavity around residues Leu-12, Phe-53, Met-90, and Ser-56 is fairly devoid of water. The second channel starts at

TABLE V Comparison of Structural E l m & for Domestic Hen Egg-White Lysorym and Baboon a-Lactalbumin" a Helix

DHEL (A) 4-15

(B) 24-36 (C) 88-99 (D) 108-115

3 10 Helix

a-LA 5- 11 (5- 16) 23-34 (25-36) 86-99 (89-103) 105- 109 (109- 1 13)

DHEL

79-84 119-124

/3 Sheet

a-LA

DHEL

a-LA

12-16 (12-18) 17-21 (19-23) 76-82 (79-85) 101-104 (105-112) 115-1 19 ( 1 19- 126)

42-60

40-43 (42-45) 47-50 (50-53)

1-2 39-40

O(A), (B), (C), and (D), a Helix A, B, C, and D. DHEL Domestic hen egg-white lysozyme; a-LA, baboon a-lactalbumin. Numbers in parentheses signify equivalent residues in domestic hen egg-white lysozyme. Based on results by Acharya et al. (1989) and by Blake et al. (1967a).

LYSOZYME AND a-LACTALBUMIN

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Ile-27, runs to Asp-88, and is partially occupied by water molecules. The channel is “blocked” by Tyr-103 (which is in the cleft region). There are corresponding cavities in hen egg-white lysozyme. T h e second cavity in the vicinity of Ser-91 is occupied by internal water molecules in egg-white lysozyme. This residue becomes Asp (residue 88) in a-lactalbumin. Due to calcium binding properties in a-lactalbumin, the locations of internal water molecules are somewhat different from those in lysozymes that do not bind calcium. 6. T h e location of the bound calcium(I1) ion was unequivocally identified (see Fig. 8). This is probably the most important feature of this work and is further discussed in Sections VI and VII (see also Table IX). It should be emphasized here that the calcium-binding fold in a-lactalbumin resembles only superficially the “EF-hand” of those calcium-binding proteins that exhibit this feature (Friedberg, 1988; see also Stuart et al., 1986). 7. In the course of their nuclear Overhauser effect (NOE) studies of a-lactalbumin, Poulsen et al. (1980), and later Koga and Berliner (1985), reported that the “hydrophobic box” region of hen eggwhite lysozyme, first noted by Blake et al. (1967a), is conserved in a-lactalbumin. This is confirmed in the X-ray crystal structure: residues Ile-95, Tyr-103, Trp-104, and Trp-60 form the box. In contrast for hen egg-white lysozyme the box is composed of residues Tyr-20( 18), Tyr-23(21), Trp-28(26), Trp-108( 104), Trp-

FIG. 8. Stereo view of the Ca(I1) binding site in baboon a-lactalbumin. (Reproduced with permission from Phillips et al., 1987.)

214

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

111(107),Leu-17(15), Ile-98(95),and Met-105(101). (The equivalent a-lactalbumin numbering is shown in parentheses.) It should be noted that Ala-107 in lysozyme is replaced by Tyr-103 in alactalbumin, with the potential of blocking saccharide binding, to which we have alluded above.

C . Conclusions In conclusion, attention is drawn to several puzzling features: the differences found in the cleft region suffice to predict that a-lactalbumin would have no cell lytic activity. It remains an anomaly, however, that weak activity has been demonstrated for a-lactalbumin from various sources by McKenzie and White (1987) (Section X), and it is an unresolved problem as to how such activity could be explained, except by the possible involvement of His-32 in a-lactalbumin as an active site residue, in place of Glu-35, which appears in lysozyme (for further discussion see Section X). In addition, there are numerous discrepancies between the reactivities of a-lactalbumin and lysozyme. The former is generally a more reactive protein (Section IX), and these differences could not have been predicted by consideration of the above models, nor from the X-ray structural analysis. BINDINGOF METALIONS IN LYSOZYME VI. COMPARATIVE AND a-LACTALBUMIN

A. Introduction We have already seen in Section V that the determination of the highresolution structure of a-lactalbumin was frustrated by a variety of problems. Eventually, the evidence for the binding of calcium in the protein crystals led the Phillips group to a structure in which the binding site is part of a specific structural feature. Proposals for the binding of calcium were heavily dependent on previous studies of calcium binding by a-lactalbumin in solution, as well as on amino acid sequence studies. It is about 10 years since the binding of calcium by a-lactalbumin was first noted, and since that time the binding has been intensively studied, resulting in a voluminous and sometimes conflicting literature. Metal ion binding by lysozyme has been studied over a somewhat longer period. A brief review of the studies of both proteins is important for comparative purposes and for elucidation of the evolutionary rela-

LYSOZYME AND (Y-LACTALBUMIN

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tionships of the two proteins. This comparison takes on a particular significance, since it constitutes an area of contrast between the two proteins, against a background of structural similarity.

B . Metal Ion Binding to Lysozyme The first systematic investigation of the binding of a metal ion by lysozyme is probably that by Fiess and Klotz (1952), who found the affinity of five proteins for Cu(I1) at pH 6.5 to be in the order bovine a-casein > &casein > serum albumin > P-lactoglobulin > hen egg-white lysozyme. Soon afterward Carr (1953), using membrane electrodes, found that -0.7 mol of Ca(I1) was bound per mole of hen egg-white lysozyme at pH 7.4, compared with 6.7 mol of Ca(I1) per mole of bovine serum albumin. In commenting on the lack of correlation of Ca(I1) binding for different proteins with their isoelectric points, Carr ( 1953) made the perceptive statement: there is a heterogeneity in available binding spots which has not been fully explained. It is most likely that the explanation lies in the structural relationships between the various active groups as they occur in a particular protein molecule. Thus a further understanding of these interactions will await further information about protein structure such as the effect of hydroxyl and other functional groups, amino acid sequences, and the three dimensional nature of the polypeptide chains.

Some years later, McDonald and Phillips (1969) studied a shift in the nuclear magnetic resonance (NMR) spectrum of hen egg-white lysozyme induced by Co(I1) and concluded that this cation participates in coordinative binding to a single site. Gallo et al. (1971), using electron paramagnetic resonance (EPR), studied the binding of Mn(II), as well as Co(II), to lysozyme. The binding of each involved Asp-52 and Glu-35. Both metal ions are inhibitors of lysozyme activity, but Mn(I1) binds more strongly than Co(I1).Jori et al. (197 1) coordinated Zn(1I) as well as Co(I1) to lysozyme and again found Glu-35 and Asp-52 to be involved. Ikeda and Hamaguchi (1973) studied the binding of Mn(II), Co(II),and Ni(I1) to lysozyme by circular dichroism (CD) and determined their binding constants. Teichberg et al. (1974) studied the binding of Cu(I1) to lysozyme by spectrofluorometry and X-ray crystallography. With spectrofluorometry, they determined that Cu(I1) was located in the neighborhood of Trp-108, the association constant being 1.8 x lo2M - I . This observation was confirmed and extended by their use of X-ray analysis, whereby Cu(I1) was placed at 0.7 nm (7 A) from Trp-108. In addition, this cation was found to be 0.2-0.3 nm (2-3 A) from the car-

216

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

boxyl side chain of Asp-52 and 0.5 nm (5 hi) from that of Glu-35. Secemski and Lienhard (1974), measuring proton release and ultraviolet (UV) difference spectra, found that Gd(1II) is also bound between these residues, being attached to their carboxyls at the junction of binding sites D and E, in accordance with X-ray crystallographic findings. Kurachi et al. (1975) made a crystallographic study of the positions of Mn(II), Co(II), and Gd(II1) in triclinic egg-white lysozyme. The first two of these were 0.25 nm (2.5 hi) from one of the oxygen atoms in the Glu-35 side chain. There were two Gd(II1) binding sites. The one of highest affinity was 0.32 nm (3.2 A) from an oxygen in the Glu-35 side chain, and the other was 0.32 nm (3.2 h;) from an oxygen in the Asp-52 side chain. Jones et al. (1974) determined the water-proton relaxation times of the Gd(II1)lysozyme complex in aqueous solution. Perkins et al. (1979), using X-ray analysis, also found that Gd(1II) binds at two sites, one close to Glu-35 and the other close to Asp-52 [cf. the lanthanide complexes (Dobson and Williams, 1977)l. The two sites are 0.036 nm (0.36 hi) apart. There were numerous small conformational changes on the binding of Gd(III), as well as NAG, which had been complexed with Gd(II1). Some 13 years after Carr’s original observations, Kretsinger (1976), in his review of calcium-binding proteins, assumed that lysozyme can attach Ca(II), as well as other cations. It was not until 1981 that binding of Ca(I1) to lysozyme was further studied. Imoto et al. (1981) determined the stability (association) constant (40 M - I ) and found that lysozyme is inhibited in the presence of Ca(II), showing only 26% of the activity of the free enzyme toward hexa-N-acetylglucosamine. Because of this inhibition, they predicted that Ca(I1) binds near the catalytic carboxyls. Furthermore, Ca(I1) shifts the native-denatured transition in lysozyme toward the native state, and thus has some preservative effect on the protein. We will see in Sections VII and X that the recent elucidation by X-ray crystallography of the binding sites for Ca(I1) in baboon a-lactalbumin has led to a flurry of studies of potential binding by variants of lysozyme in a wide range of species.

C . Metal Ion Binding to a-lactulbumin

The first substantive report of the binding of Ca(I1) by cw-lactalbumin appears to be that by Hiraoka et al. (1980), who found that there is one site to which Ca(I1) is strongly bound in this protein, and some evidence of other weak binding sites. They concluded that a-lactalbumin is a cal-

LYSOZYME AND a-LACTALBUMIN

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cium metalloprotein, and that calcium stabilizes the protein against unfolding by heat and by guanidine hydrochloride. This work led to an investigation by Permyakov et al. (1981), who studied a low-pH conformational shift involved in binding of one Ca(I1) to a-lactalbumin, which caused a change in the Trp fluorescence quantum yield and a spectral shift toward shorter wavelengths. They concluded that the shift at low pH resulted from competitive replacement of the bound Ca(I1) by hydrogen ions. On the basis of fluorescence changes during EGTA {[ethylenebis(oxyethylenenitri1o)ltetra-aceticacid}titration of Ca(I1)-a-lactalbumin and in pH titrations, Permyakov et al. (1981)found that the first association (stability) constant (&) for Ca(I1) binding by bovine a-lactalbumin is 4.5 ( & 1.5) x lo* M - I . Furthermore, Van-Ceunebroeck et al. (1985) found Ks,lfor bovine a-lactalbumin to be greater than lo7M - I . Herein lies a major difference between the 1: 1 binding of Ca(I1)by bovine a-lactalbumin and domestic hen egg white, the ratio of the two association constants being on the order of lo7:1. (The values of Ks,l determined by Kronman’s group would give a ratio of lo5: 1, for which see below.) During the past 10 years Berliner and associates have made an extensive study of the binding of metal ions by a-lactalbumin and their role in the action of lactose synthase (for review see Berliner and Johnson, 1988).This work includes a study by Murakami et al. (1982) of the binding strength of Ca(I1) by bovine, caprine, human, and guinea pig alactalbumins. They found that & for Ca(I1) is of the order of 1010-1012 M-1, and that for Mn(I1) is -lo6 M-’. They also concluded, on the basis of hypsochromic wavelength shift and quenching of Trp fluorescence, that the metal ion induced a conformational shift. As well as the strong binding site, they found evidence of three weaker binding sites. Finally, they stressed the need for determination of the equilibrium constants by a method such as ESR, in addition to the fluorescence method, in order to avoid potential errors. Soon afterward, Kronman, who has made a long study of a-lactalbumin reactions, considered that there was an experimental artifact in the use of chelating metal ion buffers (e.g., EGTA and EDTA) in the determination of association constants for metal ions with proteins by fluorescence titration. Kronman and Bratcher ( 1983)concluded that their observations explained the discrepancy between K 1for Ca(I1)and bovine a-lactalbumin reported by Kronman et al. (1981) (2.7 X lo6 M - l ) , Permyakov et al. (1981) (6.3 x lo8 M - l ) , and Murakami et al. (1982) (4 x lo9 M - I ) . Some years later a strong rebuttal was made by these groups to the criticism by Kronman and Bratcher (1983). Permyakov et al. (1987) stated that there is no valid evidence of artifact in their determinations and

-

218

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

reiterated that the value of K S , ] for , Ca(I1) and bovine a-lactalbumin, is in the range of 0.25- 1.0 X lo9 M-I. Kronman (1989) has reiterated his criticism in his recent review of metal ion binding by a-lactalbumin. In the meantime Japanese and Dutch groups also made determinations of Ks,,. Segawa and Sugai (1983) concluded that bovine, human, and caprine a-lactalbumins “prepared by ordinary methods” contain 1.1-1.3 Ca(I1) ions per protein molecule and that the removal of the calcium “destabilizes the tertiary structures in these proteins.” They concluded, on the basis of changes in CD ellipticity, that KS,] values for these proteins are, respectively, 2.5 X lo8 M - I , 3.0 x lo8 M - l , and 2.8 X lo8 M-I. Later, Hamano et al. (1986), using a calcium-sensitive electrode, determined K s s 1in , 0.06 M Tris buffer (pH 7.8-8.5) in the presence of varying concentrations of NaC1. They found Ks,, for Ca(I1) and Na(1) to be 2.2 (+0.5) x lo7 M-I and 99 (&33) M - l , respectively, at pH 8.0 and 37°C. More nearly in agreement with Bratcher and Kronman (1984) are the results of Schaer et al. (1985), who found Ks,l for Ca(I1) binding to be from 1.2 X lo6 M-I to 2.5 X lo6 M - I , depending on the means of separation of the metal ion from the protein. The wide range of values for K , , is considered again in Section XI. D. Structural Changes on Cation Binding ly a-Lactalbumin and Their Implications in Lactose S y n t h e Activity

As indicated in Section III,B, Kronman and collaborators, in their early spectroscopic and sedimentation studies of a-lactalbumin, noted changes in a-lactalbumin as the pH was lowered below -4.0. Later, Kronman et al. (1972a,b) found that the low pH form (currently called an A form) differs from the native form (called the N form) in being somewhat less compactly folded and in a number of other properties; for example, there are changes in the environment of the tryptophan residues, but with no changes in their average extent of exposure to solvent. The nature of these changes and the origin of the terms N and A are considered in Section IX. It suffices to mention at this point that the N 4 A transition can be produced by a variety of conditions. It is now believed that the transition usually involves the dissociation of Ca(I1) from the a-lactalbumin. More recently, Kronman and Bratcher (1984) found that Tb(II1) displaces Ca(1I) in a-lactalbumin. With increasing concentration, Tb(II1) binds to a second site with a concomitant decrease in affinity for metal ion binding to the first site, resulting in a decreased stability of the native conformation (or N) conformer, and thus renders more favorable the

LYSOZYME AND a-LACTALBUMIN

219

conversion of a-lactalbumin to an “A-like” state, as determined by fluorescence measurements. The term “A state” was used first by Kuwajima (1977) and appears to be synonymous with “U state,” which had been used by Kronman and co-workers (e.g., Kronman et al., 1972a,b, 1981) to denote not only the conformational state that results from acid denaturation, but also that which results from Ca(I1) removal. Much work has focused on partially folded conformers of this protein (for more discussion see Section IX,A, E, F, and H). Kronman and Bratcher (1984) found additionally that a third, weaker, binding site exists for Tb(II1) in a-lactalbumin, and concomitant with this binding was a further conformational change, as judged by fluorescence properties, which they termed the “expanded” A-like state. Kronman and Bratcher (1984) found two binding sites for Zn(I1) in bovine a-lactalbumin. At the site of lower affinity, Zn(I1) caused conversion to the expanded A-like state [presumably the same as that seen also (above) with Tb(II1) binding]. There appear to be three binding sites in a-lactalbumin for Mn(I1). Of particular interest in the study of effects of metal ion binding on the conformation of a-lactalbumin are the contributions of Berliner and co-workers. Murakami et al. (1982), in a study of bovine, caprine, human, and guinea pig a-lactalbumin, observed metal ion-induced conformational change resulting in a unique hypsochromic shift and quenching of tryptophan fluorescence. They found that Ca(I1) and the lanthanides Tb(III), Eu(III), Gd(III), Yb(III), Pr(III), and Dy(II1) could be bound extremely strongly to a specific site. They also found that Mn(II), Ca(II), and Mg(I1) could be weakly bound to the same site. Murakami and Berliner (1983) later reported the existence of a zinc binding site in bovine, human, guinea pig, and rabbit a-lactalbumins, in which the zinc site is physically distinct from the site for binding calcium. This proposal was supported by the fact that when a cation binds to one site, the ensuing conformational shift excludes binding to the other site. All metal ions that were bound to apo-a-lactalbumin at the calcium site caused the same fluorescence shift. Titration of Ca(I1) or Mn(I1) protein with Zn(I1) or Al(II1) caused a complete return to apo-a-lactalbumin fluorescence parameters. I n contrast, titration of apo-a-lactalbumin with Zn(I1) caused no change in fluorescence parameters. Berliner et al. (1983) sabstituted l13Cd(II) or Mn(I1) for Ca(1I) in bovine and caprine a-lactalbumins. On the basis of NMR and ESR studies, respectively, of the 113Cd(II)and Mn(I1) proteins, they concluded that coordination to the metal ions was through oxygen. They considered the relationship of the binding site in a-lactalbumin to the “EF-hand domain” in calcium binding sites, as discussed in Section V.

220

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

By studying the binding of the fluorescent probe, 4,4’-bis[l-(phenylamino)-8-naphthalene sulfonate] (bis-ANS), Musci and Berliner (1985a) were able to differentiate between a new apo-like conformation, which was locked in by binding with either Zn(I1) or AI(III), the true apo form with no metal ion attached, and that induced by the binding of Ca(I1). They concluded that their experimental evidence enabled a distinction to be made between site I, the calcium binding site, and site 11, the site that binds Zn(I1). T h e results also were suggestive of a-lactalbumin possessing an hydrophobic surface that becomes somewhat less accessible on 1: 1 calcium binding in the absence of metal ions that bind to site I1 [see also Desmet et al. (1987) in Section IX,A for further use of bis-ANS in the study of a-lactalbumin]. Musci and Berliner (1986), using Forster energy transfer measurements between donor Eu(1I) or Tb(II1) at site I and acceptor [Co(II)] at site 11, estimated the distance between these sites to be 11.5 _t 1.5 A. They also measured the distance between the locus of bis-ANS and Co(I1) at site I1 to be 13.6 +- 1.0 h;. Also determined was the distance between bis-ANS and a fluorescein moiety covalently bound to Met-90, which was 33.5 & 3.1 A, and between Met-90 and Co(I1) at site 11, which was 16.7 & l . O h ; . Further determinations of intramolecular distances have been made by Musci et al. (1987). Met-90 in a-lactalbumin was spin-labeled. Paramagnetic line broadening of the spin-labeled ESR lines by Gd(III), substituted at the high-affinity site, yielded a distance of 8 f 1 h; between the spin label and the metal binding site. Distances between the Met and several resolvable protons were also determined from paramagnetic line broadening, with the use of NMR. Musci and Berliner (1985b) concluded that apo-a-lactalbumin is more efficient as the modifier protein in the lactose synthase system than is the Ca(I1)-bound form. They found that V, for the apo form shows a 3.5-fold increase over that for the Ca(1I)-bound form, but there is no difference in K, (app.) between the two forms. They also confirmed that calcium stabilizes the protein against thermal denaturation (see Section IX,E), but that zinc is crucial in shifting the protein toward the apo-like form that is optimally active in lactose synthase. Their model is summarized schematically in Fig. 9. The question as to possible differences in conformation between the apo and Ca(I1)-bound forms of a-lactalbumin was also addressed by Kuwajima et al. (1986), who found that the Ca(I1)-bound and free forms can assume essentially the same folded conformation, as evidenced by similarity in their CD and proton NMR spectra. However, on the basis of CD studies of aromatic side-chain effects, they concluded that the stability of the folded state is markedly enhanced by Ca(I1).

22 1

LYSOZYME AND a-LACTALBUMIN

11

11

FIG. 9. Conformational states of a-lactalbumin in solution, as suggested by Musci and Berliner (1985b). (Reproduced with permission from Musci and Berliner, 1985b.)

In contrast to these findings are those by Van Ceunebroeck et al. (1986), who used a 1251-labeledhydrophobic dye in the study of the apo and Ca(I1)-bound forms of bovine a-lactalbumin. The former protein was more heavily labeled with the dye than the latter, and a larger hydrophobic surface was therefore concluded to be exposed in the absence of Ca(I1). Some other recent studies have been concerned with the effects of monovalent cations. Hiraoka and Sugai (1984) showed that one Na(1) ion binds to a specific site in a-lactalbumin, presumably the Ca(I1) binding site. The bound Na(1) stabilizes the native form of the protein. Hiraoka and Sugai (1985) reported that both Na(1) and K(I) stabilize the nativelike state of a-lactalbumin. However, the conformational change induced by these ions, from the partially unfolded apo form to the native form, is slow compared to that brought about by Ca(I1). Permyakov et al. (1985) studied the binding of Na(1) and K(I), as well as of Ca(I1) and Mg(II), to bovine a-lactalbumin by intrinsic protein fluorescence. Urea- and alkali-induced unfolding transitions involve stable partially unfolded intermediates for the ion-bound forms of this protein (see also Section IX,E).

222

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

E . Metal Ion Binding in a-Lactulbumin: Implications for Lysozyme The studies in solution, by several schools, of the binding of calcium to a-lactalbumin, as well as the realization that crystals of a-lactalbumin (as ordinarily prepared) contain calcium, were of great importance in the elucidation of the three-dimensional structure of baboon alactalbumin by Phillips and co-workers, as already indicated. At the same time the precise delineation of the calcium binding site, discussed in Section VII (see also Table IX and Fig. 8), naturally led to the consideration of the minimum number of these residues that must be present and the relevant conformation of the peptide chain to enable calcium binding to occur. Questions that arise immediately are: Can any Iysozyme bind Ca(I1) in a comparable manner? To what extent can lysozymes exhibit weak lactose synthase activity and a-lactalbumins exhibit weak lytic activity? A crucial issue in the binding of Ca(I1) and other metal ions to a-lactalbumin is the number and nature of the binding sites. D. C. Phillips (personal communication, 1989) only found evidence for one binding site in crystalline a-lactalbumin. In a comprehensive review of the binding of metal ions to a-lactalbumin, Kronman (1989) postulates up to six ion binding sites. This is considered in Section XI. Many years ago Hopper and McKenzie (1974) noted structural similarities between equine and echidna lysozymes. They also obtained some evidence, albeit controversial, of a weak ability of echidna lysozyme to act as a modifier in the lactose synthase system. More recently, McKenzie and White (1987) noted very weak lytic activity in a variety of a-lactalbumin preparations. Also, Teahan et al. (1986, 1990) confirmed certain essential structural features for Ca(I1) binding in echidna lysozymes I and 11 and noted the potential binding of Ca(I1) by equine and pigeon lysozymes. D. C. Shaw and R. Tellam (quoted by Godovac-Zimmermann et al., 1987) made preliminary fluorometric observations that indicated binding of Ca(I1) by echidna and equine lysozymes. Subsequently, Nitta et al. (1987) concluded that equine lysozyme was a metalloprotein, containing one Ca(I1) ion per molecule. They recently determined K , , for binding of Ca(I1) by equine and pigeon lysozymes to be 2 X lo6 M-I and 1.6 X lo7 M - l , respectively (Nitta et al., 1988). More recently Desmet et al. (1989) found that removal of Ca(I1) from equine lysozyme induces a small but significant change in CD behavior, indicating a slightly unfolded apo conformation, apparently similar to that of the apo form of a-lactalbumin. Sugai et al. (1988), Nitta and Sugai (1989), and Acharya et al. (1991) have discussed the evolution of metal binding sites in proteins. We leave

LYSOZYME AND ff-LACTALBUMIN

223

an assessment of the evolutionary aspects for discussion in Section X. In the interim we make a few other comments. Some additional comparative inference may lie in the nature of the research thus far conducted on these proteins. It is seen that, for lysozyme, emphasis has been mostly in the direction of determining the cation-binding loci for the various metal ions studied. With a-lactalbumin, on the other hand, studies of conformational changes occurring upon addition or removal of cations have been heavily emphasized in the literature, probably because such changes occur readily for alactalbumin and thus are more in evidence than for lysozyme. Hence, alactalbumin may possess greater conformational flexibility, with greater adaptability to complex formation, as is evident, for example, in its ability to combine with galactosyltransferase to form lactose synthase. That a-lactalbumin is inherently more susceptible to denaturative influences and other reactions is well established (see Section IX,E). What purpose metal ion binding may serve for lysozyme and alactalbumin in nature is largely unclear, despite proposals by some authors that cations may serve to stabilize a given conformational structure, as well as exerting control over their activities by inhibitory effects. According to Lonnerdahl and Glazier (1985), only 1% of the calcium content of human milk and 0.15% of the calcium content of cow milk are bound to a-lactalbumin. Hence, this protein is quantitatively unimportant for calcium nutrition of the infant. They point out, rather, that the primary role of calcium may be to regulate lactose synthesis and possibly to aid in the secretion of a-lactalbumin. On the other hand, Rao and Brew (1989) have found that Ca(1I) is essential for the formation of correct disulfide bonds and the development of native conformation. They suggest that Ca(I1) may function to guide the folding of the nascent protein. Musci and Berliner (1985b) have suggested that a balance between Ca(I1) and Zn(I1) may serve to “fine-tune” the protein conformation, affecting the release of this protein from binding with the membrane of the endoplasmic reticulum, as well as its modifier activity. AND SEQUENCE HOMOLOGIES IN VII. AMINOACIDCOMPOSITION LYSOZYME AND a-LACTALBUMIN

A . Amino Acid Compositions

In early comparative studies of proteins, both those of the same protein from different species and of genetic variants of a protein within a

224

H U G H A. MCKENZIE AND FREDERICK H. WHITE, JR.

given species, it was necessary to compare their amino acid compositions, because sequence information was not available. Such comparisons had limitations: At best, they enabled workers to gain some idea of the lower limit to the number of differences to be expected in an amino acid sequence (see, e.g., Cornish-Bowden, 1979). Later, the experimental determination of sequences became easier, with respect both to speed of sequencing in automated sequencers and to the improvement in sensitivity, enabling much lower amounts of protein to be sequenced. Hence, attention was naturally directed to sequence determination, but to some extent relationships of particular groups of residues in evolution may be lost from sight in the emphasis on identity of residues in individual positions. Hence, we have made comparisons of amino acid compositions of alactalbumins (Table VI), mammalian c-type lysozymes (Table VII), and egg-white lysozymes (a variety of c type and one g type) (Table VIII). Where the sequence information is available, the compositions have been deduced from these results; otherwise, the amino acid compositions are obtained from amino acid analysis of the protein. The residues have been listed in the tables in the following order: the acidic amino acids (Asp and Glu) and their amides (Asn and Gln) are listed first, followed by His and then the basic amino acids (Lys and Arg). They are followed by the remaining amino acids in, broadly, their order of increasing hydrophobicity. This order is a crude “consensus” order based on the several hydrophobicity scales discussed by Edsall and McKenzie (1983). T h e comparison of the amino acid compositions may be summarized as follows: 1. On the basis of monomer molecular weights (from sedimentation-equilibrium and sedimentation-diffusion studies, amino acid sequences and compositions), the a-lactalbumins, with one exception (rat a-lactalbumin, 140 residues, see Section VII,B), have a single chain of 123 residues and M , values of 14,000. The mammalian lysozymes have 128- 130 residues and M , values of -14,400, except echidna lysozyme, which has -125 residues. The c-type hen egg-white lysozymes have -127-131 residues, in contrast to the g type, which has -185 residues. 2. All a-lactalbumins and c-type lysozymes have eight half-cystine residues (four disulfide bridges). There have been no reports of the presence of cysteine. There has been one report of a bovine a-lactalbumin having six half-cystines (Barman, 1973). As far as we know, no further work has been done on this variant. 3. In the a-lactalbumins (with the exception of the rat) the sum of

-

-

225

LYSOZYME AND a-LACTALBUMIN

the number of Asp and Glu residues (Asp + Glu) exceeds the sum of the Lys and Arg residues (Lys Arg) by five to nine residues (mean, seven residues). This difference is reflected in their low isoelectric points of pH -4.5. In contrast, for c-type hen egg-white lysozymes Asp Glu is substantially less than Lys + Arg ( - 6 to - 11; mean, - 8), leading to high isoelectric points. The position for mammalian c-type lysozymes is more complex. Human milk, rat urine, pig stomach mucosa, and echidna milk lysozymes have differences ranging from -6 to -8 residues, with a mean of -7. These lysozymes all have high isoelectric points (pH 11). In contrast, lysozymes from bovine stomach c z , baboon milk, equine milk, and deer stomach have differences ranging from - 1 to + 3 residues, with a mean of +2. This difference is reflected, for example, in the estimated isoelectric point for bovine c p , pH 7.6 k 0.2 (experimental value pH, 7.5 k 0.1) and the low pH for optimum catalytic activity. Another feature of the lysozymes is the marked variation in LysIArg ratios. These unusual differences and their significance are discussed in Section X. 4. In the 13 a-lactalbumins listed in Table Vl, seven have His contents of three residues per molecule, four have four residues, and two have two residues. Four of the mammalian lysozymes have two His residues, the remainder varying from one to five residues. Four of the hen egg-white proteins have no His residues, the remainder varying from one to five residues. 5. The Pro content of five a-lactalbumins is two residues per molecule; the remainder (with the exception of rat) are also low in Pro (one to three residues). Most of the hen egg-white lysozymes have two Pro residues, the remaining egg white and mammalian lysozymes ranging from one to five residues (except canine spleen). 6. Two variants of a-lactalbumin, caprine and ovine, have no Met residues, indicating that this residue plays no direct role in the lactose synthase system. Of the lysozymes only baboon milk and pigeon eggwhite lysozymes have no Met residues. 7. The numbers of Tyr, Trp, and Phe show small variation in alactalbumins, but appreciably greater variation in lysozymes. The greatest of the latter variations is the absence of phenylalanine in chachalaca lysozyme (see also Section VI1,B).

+

+

-

B . Sequence Comparisons The sequence comparisons of a-lactalbumins and of mammalian and avian c-type lysozymes that have been made in this article are summarized schematically in Fig. 10. In general, as far as practicable, only those sequences that have been

TABLE VI Amino Acid Compositions of a-Lactalbuminsfrom Vanow Mammals Number of Residues per Monomer Amino acid residue Asp

+ Asn

Glu

+ Gln

His LYS

'4% Pro M cys Met Ser Thr GlY Ala

Bovine B

Caprine

Ovine

13+8 = 21 8+ 5 = 13 3 12 1 2 8 1 7 7 6 3

1 4 + 8 = 22 6+ 7 = 13 3 13 1 2 8 0 6 6 5 5

1 4 + 8 = 22 6+ 7 = 13 3 13 1 2 8 0 5 5 5 6

Porcine B 21 11

3 11 1 1 8 4 6 7 7 3

Camel

Human

1 3 + 9 = 22 10+ 4 = 14 3 13 3 1 8 3 6 5 7 3

1 2 + 4 = 16 8+ 7 = 15 2 12 1 2 8 2 8 7 6 5

Equine A

Guinea Pig

1 1 + 6 = 17 8+ 6 = 14 2 12 2 3

1 6 + 4 = 20 6+ 6 = 12 4 11 2 2

8

8

3 8

1 8 6 4 5

7 7 2

Red Red-necked kanWallaby garoo

Rabbit

Rat

1 0 + 9 = 19 9+ 5 = 14 3 12 2 3 8 2 8 10 5 2

1 2 + 5 = 17 15+ 3 = 18 3 10

2 7 8 2 9 7 8 9

1 1 + 5 = 16 9+ 8 = 17 4 9 2 3 8 2 7 4 7 6

Grey kangaroo

16

16

19

19

4 10 3 3 8 2 7 5 7 6

4 10 3 4" 8 3 7 5 7 6

Leu Val lle Tyr Trp Phe Total Methodb, Refc Other variants

.I

13 6 8 4

4 4

13 6 8 4 4 4

13 5 7 4 4 4

12 2 10 4 4 4

11 2 10 3 5 4

14

2 12 4 3 4

13 4 9 4 4 4 123

s (7)

14 3 12 5 3 3

13 4 8 2 4 3

9 10 8 4 4 5

11 5 10 3 3 4 121

I1 5 9 3 3 5

-126

11 5 9 3 3 5 -127

S (11) A (12) A ( 1 2 )

'Very approximate value. bMethods for deriving composition: S, T h e composition has been determined from the complete amino acid sequence; A, the composition has been obtained from amino acid analysis. (References: ( I ) Brew et al. (1970), Vanaman el al. (1970, Shewale et al. (1984); (2) MacGillivray et al. (1979), Shewale et al. (1984); (3)Gaye et al. (1987); (4) Bell et al. (1981~); (5) Beget al., (1985); (6) Findlay and Brew (1972), Hall etal. (1982); (7) Kaminogawa et al. (1982, 1984);(8) Brew (1972), Hall etal. (1982); (9) Hopp and Woods (1979); (10) Prasad etal. (1982); (11) Shewale etal. (1984); (12) McKenzie elal. (1983). dThree genetic variants: A, B, and C. A differs from B by substitution of Gln for Arg at position 10 (Bell et al., 1970); substitution in C is not known (Bell et al., 1981a). Minor components: see Section lI1,B. 'Minor components have been identified by Schmidt and Ebner (1972); see Section Il1,B. The amino acid analysis values of 3 for Pro in caprine and ovine [given by Schmidt and Ebner (1971)] are too high. fTwo genetic variants: A and B. A differs from B in having as its amino-terminal residue Arg instead of Lys (Bell et al., 1981c). The value for Pro in the B variant is given correctly in Table 4 of Bell et al. (1981c), but it is printed incorrectly as 1.9, instead of 1.0, in Table 3. The value for Pro in the A variant has not been determined precisely, but it is assumed to be the same as for B. ZGodovac-Zimmermann et al. (1987) have examined two variants, B and C, from equine colostrum, having three and four differences from A, respectively. hHopp and Woods (1979) showed that rabbit a-lactalhumin is a glycoprotein. 'Brown el al. (1977) found that rat a-lactalbumin contains 13.4% (w/w) carbohydrate.

TABLE VII Amino Acid Compositions of Mammalian c-Type Lysozymes Number of Residues per Monomer

N N 00

Amino acid residue Asp+Asn Glu

+ Gln

Human milk, leukemic Baboon urine milk 8+10 = 18 3 6

+

= 9 His LYS '4% Pro VZ cys Met Ser Thr GlY Ala

1 5 14

2 8

2 6

9+11 = 20 3 + 8 = 11 3 5 8 3 8

Bovine stomach mucosa c2

Deer stomach mucosa

Langur stomach mucosa

Pig stomach mucosa 3

9 + 9 = 18 3+ 9

7 + 8 = 15

7 + 8 = 15

8+ 2

9+ 3

7+11 = 18 5 + 5

12

= 10

1 0 + 9 = 19 3+ 6 = 9 2 13

Horse milk

Rat urine

10+13 = 23 6+ 2 = 8 2 15 4

12

1

4

8

8

=

=

12

2 8

10 2 9 6 3 8

2

4

3

6

11 3 2 8

10 4

=

0

4

1

1

I

0

7

13

7

13 8 8 10

10

8 5

5

6

1

6

11

10

7

14

12

11

10 11

9 10 9

I1

13

Rabbit spleen

9 + 9 = 18 3+ 4

20

18

17

11

10

13

1

1

5

6 6 5 8 2 9

8 8

5 9

Canine spleen

= 7

3 8

5 15 3 3 8

1 10 4 8 10

9 9 9 8

7

Grey kangaroo

Echidna milk 1

1

8

1

8 2 9

8

7

7

12 12

10 15

1

7 6 10 10

Leu Val Ile TYr TrP Phe Total Method," Ref Other variants

r c

(0

8 9 5 6 5 2

8 9 7 6 5

10 5 3

6 6

9 9 5 5

9 9 5 5

6

2

6 2

2

6 9 7 6 5 3

10 3 7 4 4 2

8

10 6 6

2

5 5

7 8 4 2

130

130

129

130

129

129

130

130

125

s (1)

s (2)

s (3)

s (4)

s (5)

S (6)

s (7)

S (6)

S (8)

e

f

4

C

d

5

10 6 7 3 2 3 -130 A(9)

9 9 5

3 6 3

8 8 4 4 5 3

-139

-124

A(10)

A(11)

"Methods for deriving composition: S, The composition has been determined from the complete amino acid sequence; A, the composition has been obtained from amino acid analysis. 'References: Jo1lt.s and Jolles (1971, 1972), Canfield etal. (1971), Thomsen etal. (1972); (2) Hermann etal. (1973); (3) McKenzie and Shaw (1982, 1985); (4) White etal. (1977); (5) Jollts et aE. (1984), A. C. Wilson (personal communication, 1983); (6)Jolles etal. (1989); (7) Stewart etal. (1987), includescorrection given by Jolles etal. (1989); (8) Teahan et d.(199lb); (9) Jolles and Fromageot (1954); (1O)Jolles and Ledieu (1959); McKenzie et al. (1983). [ A similar, but not identical, lysozyme has been isolated from donkey milk by Godovac-Zimmermann el al. (1988). d T ~ other o variants, c, and q ,have been identified by Dobson rt al. (1984) and by Jolles et al. (1984).
TABLE VIII Amino Acid Comflositiom of Avian Egg-White Lysozymes ~

~~~~

~~~~~~~~~~~

Number of Residues per Monomer

10 03

0

Amino acid residue Asp Glu

+ Asn

+ Gln

Black swan Domestic hen 74-14 = 21 2+ 3

C

17 8

= 5 His LYS

A% Pro Y2 cys Met Ser Thr GlY Ala

1

6 11 2 8 2 10 7 12 12

0 11 10 2 8 2 8 8 11 11

California Bobwhite quail quail

g 1 2 + 8 = 20 6 + 8 =14

5 18 11 5 4 3 9 14 21 15

7+14 = 21 2 + 2 = 4 2 6 11 2 8 2 10 7 12 12

8+13 = 21a 2+ 3 = 50

1 7 10 2 8

Turkey 7+13 =

20"

2+ 1 =

3a 2 7 10

2 8

2

2

10 7 12 12

10 7 13 13

Ringnecked pheasant 8+12 = 20 2+ 1 = 3 2 8 9 2 8 3 10 7 14 11

Guinea hen

Kaki duck 11

Peking duck 1

8+12 = 20 2 + 3

8+11

8+11 =19 3 + 2 = 5

=

5 2 8 10 2 8 2 10 7 12 12

=I9 4 + 1 = 5 0

6 13

2 8

0 6 13 2 8

2

2

11 7 12 11

11 7 12 11

Chachalaca 8+11 = 19

2 + 2 =

Japanese quail 9+13 = 22" 2+ 3

4

= 5 a

2

0 5 12

9 10 2 8 3 10

2 8

2 10

7

7

11 12

13 12

Pigeon 8+11 = 19 4+ 3 = 7 2 13 10 3 8 0 7 5 11 8

7 11 13 9 3 3

8 7 5 3 6 3

8 7 5 3 6 3

9 5 6 4 6 2

8 7 6 4 6 2

7 7 5 3 6 3

8 7 6 5 6 1

8 7 6 5 6 1

7 5 7 7 6 0

8 5 8 2 6 4

129

185

129

129

129

130

129

129

129

129

131

A (2)

S (3)

S (4)

S, P (5)

S, P ( 6 )

S (7)

S, P (8)

Leu Val Ile T Yr TrP Phe Total Method? Ref' Other variants

s (9) s, P ( 1 0 ) d

127

s (11) s, P(12)

d

'Exact distributions of Asp-Asn and of Glu-Gln are uncertain. "Methods for deriving composition: S, the composition has been determined from complete amino acid sequences; A, the composition has been determined from amino acid analysis; P, some of the residues have been obtained by peptide compositions and/or assumed identities. 'References: (1) Canfield (1963), Jolles et al. (1963), Canfield and Liu (1965), Imoto et al. (1972), Rees and Offord (1972), Phillips (1974), Ibrahimi et al. (1979; (2) Arnheim et al. (1973); (3) Simpson et al. (1980); (4) Ibrahimi et al. (1979); (5) Prager et al. (1972); (6) La Rue and Speck (1970); (7) Jollks et al. (1979a); (8)Jollks et al. (1972); (9) Hermann and Jolles (1970); Hermann etal. (1971); (10) Kondo etal. (1982); (11)Jolles etal. (1976); (12) Kaneda etal. (1969); (13) Rodriguez etal. (1985). dTwovariants of Kaki duck-namely, I1 and 111-and three variants of Peking duck-namely, 1, 2, and 3-have been studied. For their proposed relationships see Section VI.

232

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

completely determined are included in the comparisons. It has become evident, as substantial numbers of these proteins have been sequenced, that there is sufficient variation in sequence that it is unsatisfactory to designate residues (especially critical residues) on the basis of assumed identity or amino acid composition of peptides. However, in some lysozymes, as indicated below, we have had to make certain assumptions, especially in the designation of some Asp, Asn, Glu, and Gln residues. With improved methodology for sequence determination, important corrections have been made to some original sequences (as shown below). The basis of the individual sequences presented is summarized as follows (the notes are numbered to refer to the numbers in parentheses on Fig. 10, indicating the variant of the protein for which the sequence appears in Fig. 10). 1 . Bovine [Bos (Bos}] a-Lactalbumin

The sequence of 123 residues shown is that of the common B variant of Bos (Bos)primigenius f.d. taurus (f.d. = forma domestica) determined by Brew et al. (1970) and Vanaman et al. (1970). It includes the corrections made subsequently by Shewale et al. (1984): Residue Now Was

43 Gln Glu

46 Asp Gln

49 Glu Asp

82 ASP Asn

83 Asp Asn

87 Asp Asn

88 ASP Asn

The amino acid sequence derived by Vilotte et al. (1987; see also Hurley and Schuler, 1987) from the nucleotide sequence differs from the corrected chemical sequence presented here as follows: Residue Nucleotide Chemical

39 Gln Glu

63 Asp Asn

66 Asn Asp

Note (1): The A variant from Bos (Bos) namadicus f.d. indicus differs from the B variant at residue 10: Gln in A substituted for Arg in B (Bell et al., 1970). The sequence difference for the C variant from Bos (Bibos)javanicus (Table 11)is unknown.

2 . Caprine (Capra hircus} and Ovine (OvzS aries} a-Lactalbumin The sequence of 123 residues shown for caprine a-lactalbumin is that determined by MacGillivray et al. (1979) and corrected by Shewale et al. (1984): Residue Now Was

43 Gln Glu

46 Asp

49 Glu Asp

82 Asp Asn

83 Asp Asn

Kumagai et al. (1987) confirmed this sequence, except for residue 66,

233

LYSOZYME AND a-LACTALBUMIN

which they deduce from the nucleotide sequence to be Asn, not Asp. There could be an error in the chemical sequencing, it could be a different variant, or there could be an error in the reverse transcription process. Note (2): The amino acid sequence of ovine a-lactalbumin was derived by Gaye et al. (1987) from their determination of the complete nucleotide sequence of a-lactalbumin mRNA. This sequence (not shown in Fig. 10) is identical with the amino acid sequence of caprine a-lactalbumin except for residue 8 (assuming residue 66 in caprine is Asn): Caprine Ovine

Val Ala

3 . Guinea Pig (Cavia porcellus) a-Lactalbumin The original sequence of 123 residues was due to Brew (1972),but the subsequent DNA studies by Hall et al. (1982) resulted in the following important corrections [some of these residues are important in the binding of Ca(II), see Section V]: Residue Now Was

46 Asp Asn

57 Asp Asn

84 Asp Asn

82 Asp Asn

71 Asn Asp

87 Asp Asn

88 Asp Asn

102 Asp Asn

4 . Human (Homo sapiens) a-Lactalbumin

The sequence of 123 residues is that of Findlay and Brew (1972), with corrections from the DNA studies by Hall et al. (1982): Residue Now Was

45 Asn Asp

46 Glu Gln

82 Asp Asn

84 Asp Asn

87 Asp Asn

88 Asp Asn

102 Asp Asn

The X-ray crystallographic determination of the structure of alactalbumin has been made on baboon (Papio cynocephalus) a-lactalbumin. Its amino acid sequence has not been determined. However, on the basis of preliminary work by R. Greenberg (personal communication), it is evident, as would be expected, that its sequence is similar to that of human a-lactalbumin. On the basis of their X-ray studies, Acharya et al. (1989) concluded that the following are possible sequence changes from that of the human protein: Residue Human Baboon

11 Leu Asn

13 Lys Tyr

20 Gly Arg

58 Lys Ala

66 Val Ser

76 Ser Thr

105 Leu Ile

123 Leu Glu

5. Rabbit (0ryctoIagu.s cuniculus) a-Lactalbumin This sequence of 122 residues is that of Hopp and Woods (1979). It is predominantly a glycoprotein with the carbohydrate attached at residue 45 (Asn).

Key to Protein Abbreviations

Mammalian Proteins a-Lactalbumin Bovine B variant Caprine (goat) Guinea pig Human Rabbit Rat Equine (horse) N w Camel rp Red-necked wallaby Milk, urine, and stomach mucosa c-type lysozymes Human milk (and human leukemia urine) Baboon milk Equine milk Rat urine Bovine stomach mucosa, c type 2 (6) Deer stomach mucosa 1 Langur stomach mucosa Pig stomach mucosa 3 Echidna milk Tachyglossus aculeatus multiaculeatus, type I ( 8 )

Avian Proteins (BB a-la) ( 1 ) (C a-la) (2) (GP a-la) (H a-la) (Rb a-la) (Ra a-la) (E a-la) (3) (Ca a-la) (RW a-la) (HM lz) (BaM lz) (EM 12) (4) (RU 12) ( 5 ) (BSs Iz) (DS lz) (LS 12) (Piss lz) (7) (TMI lz)

c-Type lysozymes Domestic hen California quail Bobwhite quail Turkey Ring-necked pheasant Guinea hen Kaki duck I1 Peking duck 1 Chachalaca Pigeon

(DH lz) (CQ 14 (BWQ 12)

(T 12) (RNP lz) (GH lz) (KDII lz) (9) (PDl Iz) (9) (Chac lz) (P 12)

Key to notes (1)-(9) (see text). ( 1 ) Bovine a-la A (2) Ovine a-la (3) Equine a-la B,C (4)Donkey lz (5) Mouse M lz (6) Bovine stomach lz 1,3; Ovine stomach lz 1,2,3; Caprine stomach lz 1,2; Camel stomach Iz 1 (7) Pig stomach lz 1,2 (8) Echidna lz I1 (9) Kaki duck lz I; Peking duck lz 2,3

07.

T

FIG. 10. Comparison OF sequences of a-lactalbumins and lysozymes, including a key to the abbreviations. l’he highest numbers of-residues showing homology in a given position are boxed with continuous lines, the next highest are boxed in broken lines, and the third highest are boxed in dotted lines. a-la,a-Lactalbumin; lz, lysozyme. For further details see text. (Reproduced from H. A. McKenzie. Copyright 0 1983- 1989 by H. A. McKenzie.)

236

60

BB

c

o-la(1)

GP H Rb Pa

*-la121 a-la o-la o-la *-la

,z

(I-1a111

ca *-la

Ru

HM Bau EM RU BL DS LS Pis)

TUI

o-la

1Z

1r 1IaI 17.1II

lz I 0 11

1r 1~17)

17. 181

DH

lr

CQ

17.

B W T RNP GH KDII

1E

lz lZ

lz

PD1

lzlrl 1z 191

Chac P

1z 17.

FIG. 10. See legend on p. 235.

70

80

BB

c GP

11-11(11

.-la (21 0-1s

n a-la --la Pa a-la E *-la (:l ca e-1a Rb

m

.-la

Hn

17.

Bald

1Z

En

11111 lZ(11

Ru BSs 0s

1r.16) 1z

IS Pis)

1r

M I

lE(O1

DH

1Z

CQ BWQ

17.

lZ(7,

17.

T RWP

1z

GH

17.

KDII PD1 Chac

lZ(91 lz(9)

P

1r

1r

lr

90

100

239

240

HUGH A. MCKENZIE A N D FREDERICK H. WHITE, JR.

6 . Rat (Rattus nomegtcus) a-Lactalbumin Rat a-lactalbumin is unique in having a sequence of 140 residues. The sequence shown is that of Prasad et al. (1982). There are appreciable discrepancies between it and the nucleotide sequence of Dandeker and Qasba (1981). K. E. Ebner (personal communication, 1986) was unable to explain the difference between the work of his laboratory (Prasad et al., 1982) and the work at the National Institutes of Health (Dandeker and Qasba, 1981), for example: Residue Petal. DQ

38 Thr Ser

39 Glu Gln

41 Ser Ile

44 Asp Asn

59 Asp Asn

63 Glu Ser

64 Asn Ser

65 Gln Glu

67 Val Pro

102 Asn Asp

105 Leu Lys

Residues 28-58 are stated by Dandeker and Qasba (DQ) to differ only from Prasad (P) et al. at residues 39 and 44. This does not appear to be correct. Their comparison is actually made with Prasad and Ebner (1980).

7. Equine (Equus caba1lusf.d. caballus) (Perissodactyla) a-Lactalbumin The 123-residue sequence shown is that of the A variant (isolated from the milk of an Australian thoroughbred horse), as determined by Kaminogawa et al. (1982, 1984). A misprint in the 1984 paper has been corrected: Residue 10 should have read Gln, not Glu. Note (3): T h e B and C variants (isolated from the colostrum of a Persian Arab horse) differ from the A variant as follows (Godovac-Zimmermann et al., 1987): Residue A B

C

7 Glu Gln Gln

33 Ser Asn Asn

78 Asp Asn Asn

95 Ile Asp Ile

In this paper the residue at position 54 (A variant) is given incorrectly as Glu: it should read “Gln.”

8. Camel (Camelus dromedarius) a-Lactalbumin The 123-residue sequence is due to Beg et al. (1985).

9. Red-Necked Wallaby (Macropus rufogresius) a-Lactalbumin The sequence has only 121 residues and was determined by Shewale et al. (1984). 10. Human (Homo sapiens) Lysoqme The sequence of lysozyme isolated from the urine of human leukemic patients was determined by Canfield et al. (1971), and that of human milk Iysozyme was determined by Jollks and Jollks (1971). Canfield et al.

24 1

LYSOZYME AND CY-LACTALBUMIN

(1971), in their original sequence, showed a deletion after residue 100. The work by Thomsen et al. (1972) for the leukemic urine protein and that by Jolles et al. (1972) for the milk protein show that there is a ValVal sequence for residues 99-100 instead of a single Val residue. The revised sequences of both lysozymes are the same and are 130 residues long. The nucleotide sequences determined by Castafion et al. (1988), Chung et al. (1988), and Peters et al. (1989) are in accordance with the revised chemical sequence. 11. Baboon (Papio cynocephalw) Milk Lysozyme This sequence of 130 residues was determined by Hermann et al. (1973).

12. Equine (Equus caba1lusf.d. caballus) (Perissodactyla) Lysozyme The sequence (2) (Fig. 10) of 130 residues is that of McKenzie and Shaw (1982, 1985). Note (4): Recently, the sequence of donkey (Equus asinus) milk lysozyme was determined by Godovac-Zimmermann et al. (1988). They found the following differences: Residue Horse Donkey

61 Asn Ser

52 Ser Tyr

87 Glu ASP

13. Rodent Lysozymes: Rat (Rattus norvegicus) Urine Lysozyme and Mouse (Mw domesticus) Lysozyme M The 130-residue sequence of rat lysozyme, from the urine of rats bearing a transplantable chloroleukemic tumor, was determined by White et al. (1977). Note (5): A. B. White (personal communication from A. C. Wilson, 1986) compared the rat sequence with the results of a partial sequence determined by R. J. Riblet for mouse lysozyme. After making some realignments, White concluded that there were 14 changes in the 106 residues compared (residues 78-101 were not determined). Later, Cross et al. (1988) isolated and characterized both cDNA and genomic DNA of mouse (spleen) lysozyme M gene. They deduced the amino acid sequence from the nucleotide sequences, but found only 13 differences in sequence between the mouse and the rat. The differences found by both groups of workers are shown below, W and C signifying White and Cross, respectively: Rat

Mouse Worker

2 Thr Val W,C

18 Ser

Ah W.C

41

43

Gln Arg W,C

Arg Thr W,C

46 Asp Asn

W(?).C

47

Pro Arg W,C

74 Lys Val C

80 Pro

91

113

114

117

Gln

Asn W,C

W,C

Gln Arg W,C

Arg Ala W,C

Lys Asp Gln Val W(?),C W

Ah

120

122 Ser Arg

W

123 125 cly Ile Gln Val W(?),C W

126

Arg Glx W

242

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

“W(?)” signifies that the residue was identified by White only as Asx or Glx. Residue 74 was found by White to be the same as for the rat. 14. Bovine [Bos (Bos) taurus]Stomach Mucosa Lysoqme cp

Jolles et al. (1984) determined the 129-residue sequence of one of the three variants (variant 2) of the three c-type lysozymes isolated by Dobson et al. (1984) from bovine stomach mucosa. The sequence is given here. However, residue 98 has been altered from the original designation of Lys to His in accordance with the revision by P. Jolles and J. Jollks (quoted by Prager and Wilson, 1988). Note (6): Work by Joll&s et al. (1990) on the other bovine variants and two variants of caprine stomach mucosa lysozyme, and by Irwin and Wilson (1990) on three ovine variants, was in press at the time of writing and the following information was made available courtesy of Ellen Prager and Allan Wilson. Bovine stomach mucosal lysozyme c1 and c3 differ from c, as follows: Residue Bovine cq c, c3

c:,

48 Ser Gly

125 Glu

98 His Gln

Gln

The caprine and ovine stomach lysozyme variants differ from bovine as follows:

Residue Bovine cp Caprine 1 Caprine 2 Ovine 1 Ovine 2 Ovine 3

14 Lys

19 Gly Asp

37 Ser Gly

Glu Glu Glu Asp

48 Ser Gly Gly

63 Trp Phe Phe

72 Asn Asp

Gly

83 Glu

88 Asp

Ala Ala

Asn Asn Asn Asn

90 Ala Glu

98 His

128 Thr

Gln Ser Glu Glu

The 130-residue sequence of camel stomach lysozyme 1 differs from that of bovine stomach c2 in 36 positions: Residue Bovinecp Camel 1

3 Phe Trp

7 Glu Ala

11 Thr Lys

14 Lys Glu

17 Leu Met

21 Lys Arg

29 Leu Met

37 Ser Asp

63 Trp Tyr

67 Asp Asn

72 Asn His

75 Asp Asn

78 His Gly

79 Val Ser

80 Ala Asn

82 Ser Asn

87 Asn Asp

90 Ala Thr

94 Ala Gln

98 His Arg

99 Ile Val

101 Ser Arg

102 Glu Asp

-

114 Ser Asn

117 Arg Glu

118 Asp Gly

122 Ser Glu

123 Ser Gln

129 Thr Asp

(Numbering is that of camel.)

103

Pro

Lys Asp 83 Glu Val

62 Lys Arg 85 Met Leu

106 Ile Val

107 Thr Arg

41

243

LYSOZYME A N D (Y-LACTALBUMIN

15. Axis Deer (Axis axis) Stomach Mucosa Lysozyme

The 129-residue sequence of axis deer stomach mucosa lysozyme was determined by Jollts et al. (1989). There now appear to be two variants. The sequence given in Fig. 10 is that of variant 1. Irwin and Wilson (1990) have concluded that variant 2 differs from variant 1 as follows: Residue Deer 1 Deer 2

66 Asp Asn

90 Asp Ala

88 Asn Asp

94 Thr Ala

117 Gly ASP

16. Langur (Presbytis entellus) Stomach Mucosa Lysozyme

The 130-residue sequence of langur stomach lysozyme was determined by Stewart et al. (1987). 17. Pig (Sus scrofa) Stomach Mucosa Lysozyme

This protein was studied by Jol1i.s et al. (1989) in the mucosa of approximately 20 pigs. They found two variants (1 and 2) with identical mobility at pH 4.3 and a third variant (3) with a slightly higher mobility (0.97 of that of variants 1 and 2). The sequence shown is that of variant pig 3. Note (7): Differences between the three variants are: Residue Pig 3 Pig 2 Pig 1

37 Asn Asp Asp

43 Thr Ile Ile

45 Tyr His Arg

47 Pro

Val Val

50 Gln -

49 Ser -

104 Gln Leu Gln

106 Ile Val Ile

113 Lys Arg Arg

114 Ala Ala Thr

The pig 3 sequence is 130 residues long, and each of the other two variants has 128 residues because of the deletions at residues 49 and 50. 18. Echidna (Tachyglossus aculeatus) Milk Lysozymes Z and ZZ T h e sequences of echidna lysozyme I from the milk of Tachyglossus aculeatus multaaculeatus and echidna lysozyme 11 from the milk of Tachyglossus aculeatus aculeatus have been determined by Teahan et al. (1990; see also Teahan, 1986). Note (8): The sequences for variants I and I1 differ as follows: Residue I I1

13 Val Ala

37 Ser Gly

41 Ser Gln

Both sequences terminate at Cys residue 125. 19. Domestic Hen (Gallus gallus) Egg-White Lysozyme The original sequence was determined independently by Canfield (1963) and by Jolles et al. (1963). The sequences were generally in agreement, but the following differences were evident:

244

Canfield Jollits

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

40 Thr Gln

41

Gln Ala

42 Ala Thr

46 Asn Asp

48 Asp Asn

58 Ile Asn

59 Asn Ile

65 Asn Asp

66 Asp Asn

92 Val Asn

93 Asn Val

On the basis of the electron density map, Blake et al. (1965, 1967a) concluded that the correct assignment for residues 40, 41, 42, 92, and 93 is that given by Canfield (1963). In a later communication Blake et al. (1967a) concluded that residues 58 and 59 were in accord with Canfield (1963), contrary to the opinion given in their first paper. These conclusions were confirmed chemically by Rees and Offord (1972), who also showed that residues 46 and 48 were also in accord with Canfield (1963). Jolles and Jollb (1972) indicated the Canfield (1963) assignments for residues 40,41,42,58,59,92, and 93. Imoto et al. (1972) reported that J. K. Brown (personal communication, 1971) had suggested that residue 103 is Asn, not Asp, as given by Canfield (1963) and by Jolles et al. (1963). This suggestion was later confirmed by E. M. Prager (personal communication to Ibrahimi et al., 1979). T h e given sequence incorporates all of these corrections, and has 129 residues. The disulfide bridges were determined by Brown (1964), Jolles et al. (1964), and Canfield and Liu (1965), the results of the three groups being in agreement. The locations for a-lactalbumins and lysozymes are discussed below.

20. California Quail (Lophortyx californicus) Egg-White Lysozyme The 129-residue sequence is that of Ibrahimi et al. (1979). As indicated above, position 103 in domestic hen egg-white lysozyme was originally considered to be Asp, but was subsequently shown to be Asn. Unpublished experiments by E. M. Prager on the mobilities of equivalent peptides indicated that 103 is Asn in California quail and bobwhite quail lysozymes as well as for the domestic hen protein (see also Section VII,B,23 and 24). 21. Bobwhite Quail (Colinus virginianus)Egg- White Lysozyme

The 129-residue sequence was determined by Prager et al. (1972), and residue 103 is given as Asn on the same basis as that of the corresponding residue in the California quail protein (see above). 22. Turkey (Meleagns gallopavo) Egg- White Lysozyme

The 129-residue sequence determined by La Rue and Speck (1970) is the least satisfactory of the sequences given in Fig. 10, primarily because of the uncertain resolution of the nature of some of the residues presented by the authors as Asx and Glx. However, in Fig. 10 a number of

245

LYSOZYME AND ff-LACTALBUMIN

assumptions are made in allocating the residues: (1) Residues 18-19 have been given as Asp-Asn, since all c-type hen egg-white lysozymes sequenced to date have this sequence. (2) Residue 35 is given as Glu, since all other c-type lysozymes are Glu. (3) Residue 46 is assumed to be Asn, since all other c-type egg-white lysozymes are Asn. (4)Residue 52 is assumed to be Asp, since all other c-type lysozymes are Asp. (5) Residue 57 is assumed to be Gln, since all other lysozymes and alactalbumins are Gln. (6) Residue 59 is assumed to be Asn, since all other c-type lysozymes are Asn. (7) Residues 65-66 are shown as given by La Rue and Speck (1970) (i.e., Asx-Asx), but are probably Asn-Asp. (8) Residue 74 is shown as Asn, since all other c-type egg-white lysozymes are Asn. (9) Residues 77, 87, 93, 103, and 106 are shown as Asx, since there is doubt about the designation Asp or Asn. (10) Residue 119 is shown as Asp, since all other c-type lysozymes are Asp. 23. Ring-Necked Pheasant (Phasianus colchicus) Egg-White Lysozyme This sequence of 130 residues (note: the amino-terminal residue - 1 is Gly) was determined by Jolles et al. (1979a). Although they give residue 103 as Asp, strong evidence is presented on page 2747 of their paper that this residue is Asn, and it has been so assigned in Fig. 10. T h e sequence for the signal peptide of the prelysozyme of ring-necked pheasant was determined by Weisman et al. (1986) and compared with those of five other species of birds.

24. Guinea Hen (Numida meleagris) Egg-White Lysozyme The sequence of this 129-residue lysozyme was determined by Jolles et al. (1972). 25. Duck (Anas platyrhynchos) Egg-White Lysozyrne The sequence of Kaki duck lysozymes I1 and 111 (designated here KDII and KDIII, respectively) were determined by Hermann and Jolles (1970) and by Hermann et al. (1971). Those of Peking duck lysozymes 1, 2, and 3 (designated here PD1, PD2, and PD3, respectively) were determined by Kondo et al. (1982). In Fig. 10 the sequences of KDII and PDl are shown. Note (9): The differences for the variants were as follows: Residue 4 37 57 71 72 KDII Ser Ser Glu Gly Ser KDIII Glu Ser* Glu Arg Ala PD 1 Ser Ser Gln Gly Ser PD2 Ser Gly Gln Arg Ser PD3 Ser Gly Gln Arg Ser *Residue 37 for KDIII is 70% Ser, 30% Gly.

79 Pro Pro Pro Pro Arg

82 Leu Leu Leu Val Val

93 Arg Lys Arg Arg Arg

116 Arg Lys Arg Arg Arg

122 Lys Arg Lys Lys Lys

246

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

Although the revised sequence for residues 65 and 66 of domestic hen egg-white lysozyme was known when the above work was done, the old designations for these residues were given. Also, it was stated that the duck proteins had the same residues in these positions as for the domestic hen proteins. The evidence presented for the residues being Asp-Asn was tenuous. Hence, it is likely that these residues are Asn-Asp. Later, Jolks and Jollks (1984) assigned the duck residues as Asn-Asp, but gave no rationale for this. In Fig. 10 we have shown revised assignments. The assignment for residue 57 was discussed by Teahan (1986). She pointed out that KDII and PDl lysozymes have identical amino acid sequences, except for residue 57, which is given as Glu by Hermann et al. (1971) for KDIII and as Gln by Kondo et al. (1982) for PD1. Prager and Wilson (1972) have shown that both proteins have identical electrophoretic mobilities. Thus, it is likely that KDII has Gln in position 57. This is in accord with the later view of Rodriguez et al. (1987). Further, we note that all other c-type lysozymes and all a-lactalbumins have Gln in this position. Hence, we conclude that KDII and KDIII have Gln in position 57, as shown in Fig. 10. This, then, means that KDII and PD1 are identical. 26. Chachalaca (Ortalis uetula) Egg-White Lysozyme This sequence of 129 residues is that given by Jollks et al. (1976). 2 7 . Pigeon (Columba livia) Egg-White Lysozyme

The sequence of Rodriguez et al. (1985) has a number of surprising features, including termination at residue 127 (Cys),which are discussed below. C. Summary of Important Features of Comparative Sequences

1 . Chain Length

The chain lengths of all but three of the a-lactalbumins, considered in Section B, are 123 residues. One, rat a-lactalbumin, has a chain extension of 17 residues, giving 140 residues total. In their nucleotide sequence study Qasba and Safaya (1984) concluded that this extension arises from a T-to-G base change in the termination codon. Two sequences have fewer than 123 residues: Rabbit has 122 residues and red-necked wallaby has 121 residues. The majority of the mammalian lysozymes (human, baboon, and equine milk; rat urine; and camel, pig 3, and langur stomach) have 130 residues. Bovine, caprine, and deer stomach lysozymes have 129 residues. Although pig stomach lysozyme 3 has 130 residues, two of its variants (1 and 2) have 128

LYSOZYME AND a-LACTALBUMIN

247

residues. All but two avian lysozymes have 129 residues. Pigeon has 127 residues terminating at Cys. Echidna milk lysozyme, which resembles the marsupial a-lactalbumin in the Cys termination, has 125 residues.

2 . Amino-Terminal Residues T h e amino-terminal residues of the a-lactalbumins are variable, but four (guinea pig, human, equine, and camel) have Lys. This is in common with all of the c-type lysozymes except ring-necked pheasant, which has a one-residue extension (Gly); however, its next residue is Lys. This arises from a different cleavage point of the prelysozyme.

3 . Disuljide Bridges All of the a-lactalbumins have the structurally important cystine bridges in the same positions: 6-120,28-111,61-77, and 73-91. This is also the case for the equivalent positions in all c-type lysozymeshuman numbering: 6- 128, 30- 116,65-81, and 77-95; domestic hen egg-white numbering: 6- 127, 30- 115, 64-80, and 76-94. However, when the sequence of echidna lysozyme I was determined, this was no longer true (Teahan et al., 1986, 1990). There is no Cys at position 6; it occurs at position 9. The accommodation of this Cys in the structure is discussed elsewhere (Acharya et al., 1989). 4 . Invariant Residues In addition to the invariant positions of the eight half-cystine residues in a-lactalbumins, the following 27 residues are invariant: Glu-25(27), Phe-3 1(33), His-32(34), Ser-34(36), Gly-35(37), Thr-38(40), Val-42(44), Glu-49(53), Tyr-56(54), Gly-51(55), Phe-53(57), Gln-54(58), Ile-55(59), Leu-8 1(85),Asp-82(86), Asp-83(87), Asp-87(91),Asp-88(92), Lys-94(98), Ile-95(99), Gly- loo( 105),Trp- 104(log), Ala- 106(11l),His- 107(112), Leu115(120), Gln-l17(123), and Trp-1 lS(124). (The numbers in parentheses are the equivalent human lysozyme numbers.) When the sequences of pigeon and echidna lysozyme were determined, the number of residues invariant in c-type lysozymes was considerably reduced. In addition to the seven half-cystine residues that are invariant in lysozymes, the following 21 residues are also invariant (human lysozyme numbering, with hen egg-white lysozyme numbering in parentheses): Lys-l( l), Trp-28(28), Glu-35(35), Ser-36(36), Ala42(42), Asn-44(44), Ser-5 1(50), Asp-53(52), Tyr-54(53), Gly-55(54), Gln58(57), Asn-60(59),Trp-64(63), Leu-84(83), Ala-96(95), Lys-97(96), Gly105(104), Trp- 109(108), Ala- 111(1lo), Trp-l12(11l), and Asp-lPO(119). [Residue 7(7) is Glu in all except the camel.] In addition to the seven half-cystine residues, the following seven residues are invariant in all

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HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

of the a-lactalbumins and c-type lysozymes (numbering is human alactalbumin, with human milk lysozyme numbering in parentheses): Ser34(36), Tyr-50(54), Gly-51(55),Gln-54(58), Gly-100(105),Trp-104(109), and Ala-106(111).

5. Ligunds for Cu(ZZ) Binding Fallowing the establishment of a-lactalbumin as a Ca(I1)-bindingprotein and the revision of the sequences of several a-lactalbumins (see above), Shewale et al. (1984) suggested that residues 82, 83, 87, and 88 were probably ligands for the calcium. In fact, the X-ray crystallographic studies of Phillips’ group showed that three of the predicted residues (82, 87, and 88) were involved. The Ca(I1) proved to be seven coordinates in baboon a-lactalbumin: In addition to the involvement of the residues shown in Table IX (see also Fig. 8), two water molecules are coordinated. The majority of the c-type lysozymes so far sequenced do not have the residues necessary for the coordination of Ca(I1) (Table TABLE IX Restdues Relevant to Binding of Calcaum a

Residue Binding through

79(83)a C==O

84(88)

(C=O)

87(91) (COO)

88(92) (COO)

Equivalent residues in some a-lactalbumins and lysozymes

Source a-Lactalbumins* Baboonc Humanc BovineC EquineC Rabbit Red-necked wallaby Lysozymesd Human milk Horse milkc Echidna milkC Bovine stomach c2 Domestic hen egg Pigeonc

82(86) (COO)

79 LYS LYS LYS LY s Asn LYS 83 Ala LYS LYS Glu Ala LYS

82 ASP ASP ASP ASP ASP ASP 86 Gln ASP ASP Glu Ser ASP

84 ASP ASP ASP ASP Asn ASP 88 Asn Asn ASP ASP ASP Asn

87 ASP ASP ASP ASP ASP ASP 91 ASP ASP ASP LYS Ala ASP

88 ASP ASP ASP ASP ASP ASP 92 Ala ASP ASP Ala Ser ASP

Number in parentheses is the human lysozyme equivalent number. “Residues identified by X-ray crystallography as ligands for Ca(I1) in baboon a-lactalbumin (Stuart et al., 1986; Acharya et al., 1989) *All a-lactalbumin residues are numbered according to human a-lactalbumin. cKnown to bind Ca(I1). dAll lysozyme residues are numbered according to the equivalent human lysozyme numbers.

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IX). However, the establishment of the sequence of equine lysozyme, and soon afterward pigeon and echidna lysozymes, indicated the possibility that these three lysozymes bind Ca(I1). This was confirmed for the equine and echidna proteins in Canberra, and for the equine and pigeon proteins in Sapporo. However, the echidna lysozyme sequence is the only one in which all residues are identical with the equivalent residues in baboon a-lactalbumin. This matter is discussed further in Section X.

6 . Basic and Acidic Groups The comparison of basic and acidic groups and the ratio of Lys/Arg residues are of importance in discussing the work by Prager, Wilson, and colleagues on the divergence of gastric mucosal lysozymes. Relevant information from Fig. 10 is summarized in Table X and discussed in Section X.

7. Residues in Catalysis We have seen already, in the discussion of various structural approaches to the mechanism of catalysis by hen egg white lysozyme of hydrolytic cleavages, that residues Glu-35(35), and Asp-52(53, human numbering) are considered crucial. These residues are conserved in all c-type lysozymes sequenced to date. Other residues implicated that appear to be invariant are: Trp-63(64), Asn-59(60), Gln-57(58), Asn44(44), and Trp-108( 109). Other residues implicated are not invariant: Asn-103(104), Asp-101(102), Trp-62(63), Ala-107(108), Asn-46(46), Val109(110), Phe-34(34), Asn-37(37), and Arg-114(115).

8. Residues in Galactosyltransferase Interaction Residues that have been implicated in a-lactalbumin-galactosyltransferase interaction do not all occur in equivalent positions in any lysozyme. The nearest similarity is echidna lysozyme: a-Lactalbumin 31 32 35 115 117 118

Phe His G'Y Leu Gln Trp

Echidna lysozyme Equivalent 33 34 37 119 122 123

Phe His Gly Asp Lys Phe

The Ca(1I) potential binding residues are identical for echidna lysozyme and bovine a-lactalbumin (see above).

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VIII. GALACTOSYLTRANSFERASE AND THE LACTOSE SYNTHASE SYSTEM A . Gahctosyltramferases: Occurrence, Function, and Isolation 1 . Introduction There has been much interest in recent years in various aspects of galactosyltransferases, especially with regard to their possible use as markers in malignancy and their involvement with cell surface phenomena. We have already seen in Section II,B that lactose is synthesized in the mammary gland through an enzyme, lactose synthase, which consists of two main components: galactosyltransferase and a-lactalbumin. T h e latter acts as a specifier protein modifying the action of the galactosyltransferase. a-Lactalbumin and galactosyltransferase are under hormonal control in the mammary gland during pregnancy (Woods et al., 1977; Ono and Oka, 1980; Turkington et al., 1968). However, unlike a-lactalbumin, which is found only in milk, galactosyltransferase is widely distributed. MorrC et al. ( 1969), for example, studied galactosyltransferase activity in the Golgi fraction of rat liver. Schachter et al. (1970) investigated galactosyltransferase in the Golgi apparatus as it functions in the glycosylation of proteins. Cunningham et al. (197 1) found it in rat seminiferous tubules. Powell and Brew (1974a) demonstrated the presence of galactosyltransferase and glycosyltransferase in the Golgi membrane of onion stem. T h e former showed many similarities to the animal enzyme. It was manganese dependent and gave the same reaction (apparently) as the animal enzyme. It had a K , similar to that of N-acetylglucosamine (5.2 mM) and could be activated by bovine a-lactalbumin. 2 . Galactosyltransferase

Galactosyltransferase is only one member within the broader classification of glycosyltransferases that have been found on cell surfaces; these are involved in contact-mediated cell interactions in general, but more specifically are involved in fertilization, morphogenesis, hemostasis, and cell migration. This topic has been well reviewed by Pierce et al. (1980).

3 . Galactosyltransferaseas a Marker in Malignancy Numerous workers have reported elevations in the total glycosyltransferase level and in the sera and tissues of cancer patients (for a review see Weiser and Wilson, 1987). Podolsky and Weiser (1975),in particular,

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25 1

reported the total activity to be higher than normal in malignancy. These workers then isolated a cancer-associated galactosyltransferase I1 (GTII) isoenzyme and characterized it with a molecular weight of 76,000, contrary to the more usual 40,000-44,000 found in milk (Trayer and Hill, 1971). Patients with widespread metastases have elevated GTII levels (Podolsky et al., 1981). Also, of nine patients with Duke’s B lesions, seven had detectable GTII, and in each case it became undetectable after the patients had undergone curative colectomy. T h e authors concluded that GTII was a good marker for malignancy in general, and for pancreatic carcinoma in particular. 4. Present Scope In this review we are concerned primarily with one galactosyltransferase: UDPgalactose: N-acetylglucosamine 4/3-~-galactosyltransferase.For further discussion, see pp. 179 and 180. For a comprehensive review of the galactosyltransferases in general, see Ram and Munjal ( 1985).]Within the lactating mammary gland, galactosyltransferase becomes part of the lactose synthase system and is regulated by the flow of a-lactalbumin through the lumen of the Golgi apparatus (Brew, 1969). Most of the remainder of this section therefore deals with mammary enzyme. 5. Preparation of Galactosyltransferase In the original work on galactosyltransferase, it was isolated by classical procedures of salt fractionation and column chromatography (see the review by Brew and Hill, 1975). Grunwald et al. (1982) have warned strongly against the use of pH <5.0 in any steps involving the separation of casein; otherwise, loss of stability of galactosyltransferase results. An important development in the purification of galactosyltransferase has been the use of either a-lactalbumin or a substrate, bound to an immobile inert substance in an affinity column. With a-lactalbumin, either glucose or N-acetylglucosamine is included in the buffer to enhance the binding of galactosyltransferase to a-lactalbumin. The galactosyltransferase is then eluted with buffer not containing saccharide. Andrews (1970) and Trayer and Hill (1971) were apparently the first to use a-lactalbumin, bound to Sepharose in an affinity column, for the purification of galactosyltransferase. Trayer et al. ( 1974) used a-lactalbumin-agarose as a specific adsorbent for galactosyltransferase. Also for use in affinity chromatography of galactosyltransferase, several ligands structurally related to galactosyltransferase substrates have been immobilized onto CNBr-activated agarose. These include UDP-hexanolamine, N-acetylglucosamine, and galactosyl pyrophosphate (Barker et al., 1972). Of these media, probably N-acetylglucosamineagarose is the best known.

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A major problem in the isolation of galactosyltransferase has been molecular weight heterogeneity of the final product, apparently resulting from proteolysis. Klee and Klee (1972) used the protease inhibitor phenylmethylsulfonylfluoride, but they obtained mainly a low molecular weight form of galactosyltransferase. According to Magee et al. (1976), however, this inhibitor, as used, would give only 25-40% inhibition and is not sufficiently soluble to produce the desired inhibition. The latter authors preferred to use .s-amino-n-caproic acid, together with low tern perature. Kaminogawa et al. (1972) studied the milk protease, which is presumably responsible for the heterogeneity of galactosyltransferase, in comparison with plasmin, and they reported that these two enzymes may be the same. Subsequently, this proposal was confirmed (see Andrews, 1983). Powell and Brew (1974b) concluded that galactosyltransferase occurs in mature milk as a proteolytically degraded form of the galactosyltransferase that first appears in colostrum. Magee et al. (1973) found two different molecular weights for milk galactosyltransferase: 55,000-59,000 and 42,000-44,000. The effect of trypsin on the higher molecular weight form resulted in a product resembling the lower molecular weight form. They reasoned that a trypsinlike protease could therefore be acting to produce the latter in milk. This group further studied the heterogeneity of galactosyltransferase from cow milk, finding multiple forms of this enzyme (Magee et al., 1976). Prieels et al. (1975) found three forms of galactosyltransferase in human milk with molecular weights of 38,000, 43,000, and 50,000. The activity differences found among these forms suggested that conformation at the site of association between galactosyltransferase and the acceptor saccharide had been changed, presumably by enzymatic hydrolysis. In more recent years various factors affecting the stability of galactosyltransferase have been studied. Fraser and Mookerjea (1976) found the use of Triton X-100 to be helpful in stabilizing the enzyme during isolation. Fraser et al. (1980) found that it is also stabilized by albumin and by lysolecithin. Mitranic and Moscarello (1980) studied the effects of lipids on the activity of galactosyltransferase. Thus, phosphatidylcholine, phosphatidylethanolamine, and phosphatidylglycerol stimulated activity, while phosphatidic acid and phosphatidylserine inhibited activity. These observations are particularly relevant to the activity of galactosyltransferase in milk, with its lipid environment. Mitranic and Moscarello (1983) found that bovine serum albumin, as well as some other proteins, has a pronounced activating effect on the enzyme and cautioned against its use in the isolation of galactosyltransferase.

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B . Relationships of Structure to Function in Galactosyltransferase

There is at least one SH group in galactosyltransferase (Babad and Hassid, 1964; Magee and Ebner, 1974; Kitchen and Andrews, 1974). According to O’Keeffe et al. (1980a), an SH (one of three reactive SH groups) is at or near the active site. This point was considered further by Wong and Wong (1984), who studied the inactivation of galactosyltransferase from cow milk by SH-specific reagents. The SH group was found to be located in a nonpolar environment and in a region of nonrestricted rotation. It was found not to be located at the active site, as had been suggested by O’Keeffe et al. (1980a), nor at the proteinprotein interaction site between galactosyltransferase and a-lactalbumin. Various other studies, involving specific alterations of amino acid residues, have been conducted on galactosyltransferase. The results of Powell and Brew (1976~)indicate the involvement of a Lys residue in the activity of galactosyltransferase. Thus, an affinity label for the UDP binding site was created by periodate cleavage of the ribose moiety of UDP. The derivative caused inactivation, which was reversible by nitrogenous bases or stabilized by KBH, reduction. These observations were consistent with their hypothesis that a Schiff base had formed between an aldehyde of the affinity label and the amino group of a Lys residue. Evidence indicating involvement of a Trp residue in the activity of galactosyltransferase was found by Clymer et al. (1976). Thus, UV irradiation of the protein resulted in loss of one Trp with concomitant inactivation. Silvia and Ebner (1980) found Tyr to be essential for activity, since iodination with ICl caused inactivation. However, it was not certain from kinetic studies whether 2 mol of ICl was involved at one site, or one of each at two different sites. Iodination was explored further by Chandler et al. (1980) with lactoperoxidase, which catalyzes iodination of Tyr with I-. Further, they found that galactosyltransferase was inactivated by N-acetylimidazole. This observation was consistent with an essential role of Tyr for activity. However, this reagent also affects the amino groups of Lys residues. Differentiation between these possibilities is afforded by the fact that deacetylation with hydroxylamine will occur for the Tyr derivative, but not for the Lys derivative. Since only about 40% of the activity could be recovered by deacetylation, Lys could also have been involved. Note that this indication is the second for the implication of a Lys residue with galactosyltransferase activity (see Powell and Brew, 1976~). Conformational changes have been observed or suspected for galactosyltransferase. Magee and Ebner (1974) speculated on the occurrence

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of a conformational change with the addition of substrates or substrate analogs to the enzyme. They found that such ligands protected against the sulfhydryl reagents p-chloromercuribenzoate and N-ethylmaleimide, as well as against the inactivating action of trypsin, and a protective conformational change appeared to be the most reasonable explanation. Andree and Berliner (1980), using ESR and NMR in a study of the nature of the binding of Mn(I1) and UDP-galactose to galactosyltransferase, proposed a model consistent with their observed proton relaxation rates. Thus, there is a slow conformational interconversion of an initially formed rapidly exchanging conformation of the ternary complex to a second form that contributes negligibly to the relaxation. In addition, their evidence clearly indicated the presence of two Mn(I1) per molecule of enzyme (see also Powell and Brew, 1976a), and they noted that the affinity of galactosyltransferase for this ion is much higher in the presence of UDP-galactose. On the other hand, Klee and Klee (1970) found no compelling evidence that a conformational change from complexing of galactosyltransferase with a-lactalbumin could be responsible for the altered catalytic activity, which they suggested might be due to the “mass action effect” of complex formation (see also Bell et al., 1976). Powell and Brew (1976a) have investigated the effects of various metal ions [Zn(II), Cd(II), Fe(II), Pr(III), and Ca(II)] in combination with galactosyltransferase. All of these can substitute for Mn(II), although they do not produce as much activity as does Mn(I1). Two binding sites were distinguished. First, there is a “tight” binding site, from which Ca(I1) is excluded. Second, there is a “looser”binding site which may bind either Mn(I1) or Ca(I1). Mn(I1) was found to have a specific synergistic effect on UDP-galactose binding. O’Keeffe et al. (1980a) gained further insight into the nature of the metal binding sites of galactosyltransferase. The results from a variety of kinetic, spectroscopic, and affinity chromatography studies suggest specific functions for these sites. Thus, site I appears to be concerned with maintaining structural integrity, either of the active site region or of the protein as a whole, while site I1 is more closely associated with binding of UDP-galactose. As for the metal binding capabilities of these sites, Mn(I1) must be first liganded to site I, prior to a second Mn(II), as well as prior to substrate. Both sites can bind a number of metal ions. However, Ca(I1) and Eu(II1) bind only to site 11. Further, these workers have explored the topography of the galactosyltransferase molecule with fluorescence energy transfer measurements. Thus, transfer between Co(II), bound to site I, and Eu(III), bound to site 11, indicates a distance of 18 f 3 hi between them. An SH group (one of three) in galactosyltransferase was then modified with S-

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mercuric-N-dansylcysteine. Transfer measurements between the fluorescent ligand to Co(I1) in site I1 indicated a distance of 19 k 3 A. O’Keeffe et al. (1980b) also studied the a-lactalbumin-galactosyltransferase complex (see below).

C . Interactions of Galactosyltransferase and a-Lactalbumin in the Lactose Synthase System Klee and Klee (1970) found that the A protein of lactose synthase catalyzes lactose production, even in the absence of a-lactalbumin, albeit poorly, because of the high K , for glucose. a-Lactalbumin, complexed with galactosyltransferase, drastically lowers the K,,, values for glucose and N-acetylglucosamine. Depending on the substrate concentration, a-lactalbumin can stimulate or inhibit disaccharide formation, with both N-acetylglucosamine and glucose. The affinities of the two sugars are such that, under the usual conditions of activity determination, the concentration of glucose is optimal for lactose synthesis, whereas that of N-acetylglucosamine is inhibitory. These findings help to differentiate between possible ways in which a-lactalbumin could influence the production of lactose. One of these, which, on the surface, appears plausible, is that a-lactalbumin might accept N-acetyllactosamine as a substrate, this product having arisen from the enzymatic action of galactosyltransferase. The ensuing transglycosylation, whereby lactose would be produced, could then account for the effect of a-lactalbumin in the presence of galactosyltransferase. Brew et al. (1968) showed, however, that a-lactalbumin has no affinity for N-acetyllactosamineand thus could not be involved in this reaction. An alternative possibility involves an induced conformational change in a-lactalbumin as a result of complexing with galactosyltransferase,so as to produce an affinity for N-acetyllactosamine. Although this explanation appears never to have been ruled out, there would be no need to invoke it, since it is already known (Klee and Klee, 1970,1972; Andrews, 1969; Fitzgerald et al., 1970a) that galactosyltransferase has the innate ability to carry out this reaction, even in the total absence of a-lactalbumin. Therefore, the simplest explanation for the action of a-lactalbumin and galactosyltransferase, as suggested by Browne et al. (1969), is that a-lactalbumin, in complexing with galactosyltransferase, modifies the conformational structure of the latter to produce a form of the enzyme that more readily catalyzes the production of lactose, and then does so in preference to N-acetyllactosamine. According to Fitzgerald et al. (1970a,b),lactose may be synthesized at

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maximal rates by galactosyltransferase in the absence of a-lactalbumin, but in the presence of high concentrations of glucose. It was suggested that the lowering of K , for glucose, to permit maximal synthesis of lactose, is the physiological role of a-lactalbumin. In addition, however, according to Schanbacher and Ebner (1970), a-lactalbumin inhibits the transfer of galactose to N-acetylglucosamine. It does not appreciably inhibit transfer to various oligomers tested. A considerable effort has been expended, in several laboratories, on the kinetics of interactions involving galactosyltransferase. The results of Morrison and Ebner (1971a) were not in accord with any mechanism whereby substrates might add in any order, either wholly or partially random. The simplest mechanism, consistent with their kinetic data, was an ordered mechanism, whereby additions to galactosyltransferase proceed in the order Mn(II), UDP-galactose, and N-acetylglucosamine. Morrison and Ebner (197lb) concluded that additions also proceed in an ordered manner when a-lactalbumin is included, and the acceptor is glucose instead of N-acetylglucosamine. Thus, they appeared to add in the order Mn(II), UDP-galactose, glucose, and a-lactalbumin. They regarded a-lactalbumin as a special type of modifier that combines with the enzyme only after the addition of carbohydrate. Also, they found an ordered release of lactose and UDP-galactose and postulated that Mn(I1) does not dissociate during the reaction. Morrison and Ebner (1971~)were concerned with the kinetic effects of a-lactalbumin with either glucose or N-acetylglucosamine. Thus, alactalbumin appears to cause the following: (1) alternate pathways, (2) reductions in K, for either substrate, (3) a decrease in velocity with the N-acetylglucosamine reaction, and (4) an increase in velocity with the glucose reaction. Khatra et al. (1974) studied, by steady-state kinetics, the reactions catalyzed by human milk galactosyltransferase. Whether in the presence or absence of a-lactalbumin, they concluded that the reactants added in the order Mn(II), UDP-galactose, and monosaccharide. They felt, however, that their kinetic results could best be explained with a mechanism whereby a-lactalbumin attaches to the enzyme immediately before the monosaccharide, contrary to the finding by Morrison and Ebner ( 1971b). The attachment of Mn(I1) at site I in galactosyltransferase is essential for interaction with a ligand protein (e.g., ovalbumin) or with alactalbumin (Powell and Brew, 1976a).Also, according to these workers, the presence of saturating concentrations of UDP-galactose potentiates the binding of a-lactalbumin at high Mn(I1) concentration. When the ligand is ovalbumin, the binding of this protein and a-lactalbumin is mutually exclusive.

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Powell and Brew (1976b) found that monosaccharides enhance the binding of a-lactalbumin to galactosyltransferase, which is already saturated with Mn(I1). However, the binding of a-lactalbumin and monosaccharide is random and strongly synergistic. Thus, they felt that the effects of a-lactalbumin on disaccharide synthesis are more satisfactorily explained by random synergistic binding than by the ordered binding proposed by Morrison and Ebner (1971a,b) and by Khatra et al. (1974). Bell et al. (1976) also found that lactose synthesis proceeds after a random equilibrium addition of substrates and a-lactalbumin, since the initial rate parameters obtained with bovine galactosyltransferase at saturating concentrations of Mn(II), and a variety of acceptors, were inconsistent with an ordered addition. They concluded that the large decrease in K , for glucose in the presence of a-lactalbumin is primarily the result of the high degree of synergism in the binding of a-lactalbumin and glucose to the enzyme-Mn(I1) complex. Prieels et al. (1976) studied the binding of glycoconjugates with galactosyltransferase in both the presence and absence of a-lactalbumin. They concluded that, in contrast to its ability to inhibit N-acetyllactosamine production, a-lactalbumin does not inhibit the transfer of Dgalactose to oligomers of N-acetylglucosamine or to glycopeptides. O’Keeffe et al. ( 1980b) studied the galactosyltransferase-cr-lactalbumin complex after dansylation of Glu- 1 of a-lactalbumin. Resonance energy transfer measurements, with cobalt (bound to galactosyltransferase at site I) as the energy acceptor, indicated a distance of 32 A between the dansyl group and the cobalt. Since Glu-1 is close to the cleft region, this observation made it unlikely that this region of a-lactalbumin is involved in acceptor substrate binding. These authors presented a schematic model of the active site of galactosyltransferase and its interaction with a-lactalbumin, summing up their experimental results (O’Keeffe et al., 1980a,b).

D . Structural Requirements of Substrate Andree and Berliner (1978) reported that UDP-glucose was marginally active as a donor substrate for both the transferase and the synthase. Then Berliner and Robinson ( 1982) determined the structural requirements for the donor pyranose moiety of UDP-galactose in either galactosyltransferase or lactose synthase. Thus, an axial 4” hydroxyl group on the pyranosyl moiety is essential for precise substrate alignment, as is an equatorial 6 CH,OH moiety. Where one or the other is lacking, the maximal rate of glycosyl transfer is about 0.05% of that of UDPgalactose. Berliner et al. (1984) further studied the acceptor for lactose synthase.

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They found that the basic requirements are: (1) a pyranose, thiopyranose, or inositol ring structure; (2) equatorial substituents (if any) at C-2, C-3, C-4, and C-5; (3) the aglycone (at C-1) may be either a or P, although a is preferred; (4) in the absence of a-lactalbumin, galactosyltransferase will accept long-chain 2-N-acyl substituents on the glucosamine structure; (5) an equatorial amino or N-acyl substituent (e.g., mannosamine or acetylmannosamine) is also a suitable acceptor in the absence of a-lactalbumin, since both N-acetylglucosamine and N-acetylmuramic acid have complementary binding loci for the N-acyl moiety; and (6) the aglycone moiety must be equatorial ( P configuration). E . Final Remarks

Much has been accomplished, especially in recent years, toward the goal of elucidating the active sites of galactosyltransferase and a-lactalbumin. To this end, alteration of specific residues with observations of consequent effects on structure and activity is enlightening, as are metal ion effects. Where the substrate is concerned, we now have some detailed structural information for both galactosyltransferase and the lactose synthase system. The effect of a-lactalbumin on galactosyltransferase is possibly a mass action effect, culminating in the preference, under normal conditions of activity determination, for glucose over N-acetylglucosamine. Of the conformational changes that occur in galactosyltransferase, particularly concomitant with the addition of Mn(II), much remains to be learned in correlating structural changes with the onset of enzymatic activity. Finally, the weight of evidence from kinetic studies appears to support the point of view that the addition of Mn(II), glucose, and a-lactalbumin to galactosyltransferase, prior to the expression of enzymatic activity, is governed by random synergistic binding with galactosyltransferase. Galactosyltransferase,as was also shown, is a ubiquitous enzyme, while a-lactalbumin is restricted to the mammary gland, and expression of the latter is under hormonal control. The question can be raised as to whether other galactosyltransferases, from their many extramammary sources, might also react with a-lactalbumin or similar proteins, which might result in modifying enzymatic function. This possibility, in fact, occurred to Hamilton (198 1) after he had isolated a galactosyltransferase from the reproductive tract of the male rat (Hamilton, 1980). He then isolated and characterized an “a-lactalbumin-like” protein which, however, differed from its mammary counterpart. Thus, it catalyzed transfer of galactose equally to inositol or glucose, whereas only glucose

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can act as the acceptor in the mammary gland system. We suggest that a continued search for, and characterization of, such a-lactalbumin-like proteins could yield results of considerable genetic and evolutionary impact (see also Section XI). PHYSICAL, CHEMICAL, AND BIOLOGICAL IX. SOMEADDITIONAL COMPARISONS BETWEEN LYSOZYME AND ~-LACTALBUMIN

A . Spectroscopic Studies

Many of the following studies, particularly those that measure changes by absorption or fluorescence, were made for a-lactalbumin, without corresponding studies on lysozyme that may have been used comparatively. One must consider the likelihood that some such studies have been attempted, but without success, since unsuccessful experiments seldom find their way into the literature. It can only be surmised that the conformational structure of lysozyme is sufficiently more resistant to change that such studies would have proved relatively unproductive. It is well established, in fact, that lysozyme does offer more resistance to denaturative change and to chemical alteration in general (Section IX,E and F). 1 . UV Absorption Spectroscopy Kronman et al. (1965) and Kronman and Holmes (1965) appear to be the first to have studied the effects of acid on a-lactalbumin and report that this protein, adjusted to pH values below its isoelectric point, exhibits a hypsochromic shift in its absorption spectrum between 270 and 300 nm. Spectral shifts in this region usually reflect changes in the environment of Trp and Tyr residues. The conformational change is a complex one, involving a series of steps. Because of the nature of the shift, the numbers of Trp and Tyr residues present, and the relative magnitudes of E for Trp and Tyr, Kronman and co-workers concluded that the shift results from environmental alterations for more than one of the buried Trp residues. (At the time of this study, three Trp residues were considered to be buried in bovine a-lactalbumin.) There appears to be no corresponding effect for hen egg-white lysozyme. [However, note the effect of acetic acid studied by Kato et al. (1984).] 2 . UV Difference Spectroscopy

Conventional UV difference spectroscopy and solvent perturbation difference spectroscopy have been used in a wide variety of protein stud-

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ies (McKenzie, 1970; Nicola and Leach, 1976). Both methods have been used in studies on the involvement of Trp residues in various reactions of a-lactalbumin and lysozyme. Efforts with this methodology have been limited, probably because, as pointed out by Imoto et al. (1975), “even a relatively simple-appearing pH difference spectrum reflects a complex set of interactions involving several chromophores and several ionizable groups.” Hayashi et al. (1964) used both methods to study the changes that occur upon formation of the lysozyme-substrate complex. Their results indicated that at least one Trp residue becomes buried in the hydrophobic region produced by formation of the complex. Imoto et al. (1975) used difference spectroscopy to investigate saccharide binding of lysozyme and concluded that Trp-108 makes a principal contribution to this reaction. An early investigation of a-lactalbumin with the solvent perturbation method was undertaken by Kronman and co-workers, who studied the degree of exposure of Trp groups under a variety of conditions, especially the changes at low pH (-4) (Kronman et al., 1965; Kronman and Holmes, 1965; Robbins et al., 1967). When this work was initiated, it was not known that a-lactalbumin is a Ca(I1)-requiring protein (see Sections II1,B and VI,D). As already indicated, the normal form of the protein is now referred to as the N form, and the low-pH form, from which Ca(I1) is lost, is called the A form. Molecular states similar to the low-pH A form may be produced by a variety of other treatments (see Section IX,E). [Molecular states similar to the low-pH A form also may be produced by a variety of other treatments (see Kronman, 1989).] In their initial studies it was thought that the low pH conformational shift in the N + A transition involved the exposure of the buried Trp residues. However, their solvent perturbation difference spectral studies enabled them to conclude that the change at 25°C does not involve unfolding of the molecule in the region of the buried Trp groups and their consequent exposure, but involves alterations in the interactions of these Trp residues with other perturbing groups. At pH 6 and 1°C there is a loss of accessibility of the two exposed Trp groups to sucrose, but they are still accessible to water. In the pH range 1.8-3.0 at 1”C, one exposed Trp group becomes inaccessible to sucrose, but the other is accessible to water and sucrose. Thus, Kronman and co-workers visualized the exposed groups as lying in a “crevice” with contraction that occurs with decrease in temperature, the extent of which was pH dependent. Later, Kronman et al. (1972a) considered that the “crevice contraction” phenomena are probably artifactual. Kita et al. (1976) studied the reversible unfolding of a-lactalbumin in

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guanidine hydrochloride, using difference spectroscopy and pH -jump measurements. They concluded that a-lactalbumin in the native state is less stable than lysozyme, an observation that is in agreement with conclusions from many other studies (see Section IX,D-F).

3 . Fluorescence Spectroscopy Kronman et al. (1971) used both fluorescence spectroscopy and CD in a study of conformational changes occurring upon acylation of alactalbumin. The observed changes in fluorescence were similar to those occurring on acid denaturation. Thus, it appeared likely that the results of acylation, as well as acid and alkaline denaturation, are brought about by conformational changes that give rise to freedom of rotation of Tyr and Trp side chains. Rawitch (1972) reported a difference in the rate of rotational diffusion, determined from the polarization of fluorescence, which suggested that the effective molecular volume of a-lactalbumin in solution is larger than for lysozyme, and this difference suggests conformational differences. Miller and King (1975) studied the differing luminescence properties of lysozyme and a-lactalbumin. For the latter the spectral properties observed were attributed to the proximity of Trp residues to S-S bonds, the reduction of which caused much change, as did acid denaturation. Their work supports the earlier suggestion by Sommers et al. (1973), that energy is transferred from Trp- 109 to Trp-63 in the active site cleft, with subsequent quenching of the latter by neighboring cystine residues. Sommers and Kronman (1980) made further progress in the characterization of Trp chromophores in comparative fluorescence studies of bovine, goat, human, and guinea pig a-lactalbumins by characterization of the environments of individual Trp residues in partially unfolded conformers. In the native state Trp-28 and Trp-109 transfer their excitation energy to Trp-63, whose fluorescence, as suggested earlier by Kronman’s group, is quenched by a pair of vicinal S-S bridges. Changes in fluorescence occurring upon formation of the heat-denatured form are caused by exposure of Trp-63. T h e effect of low pH on a-lactalbumin was studied further by Permyakov et al. (1981). In addition to the spectral shift toward shorter wavelengths (observed by Kronman et al., 1965), they found a decrease in Trp fluorescence quantum yield. They suggested that the replacement of Ca(I1) by H(1) is basically responsible for these effects. Much more recently, Desmet at al. (1987) studied the hydrophobicity of the partially unfolded conformer of a-lactalbumin, which results from removal of Ca(II), in comparison with the native conformer, mak-

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ing use of the hydrophobic fluorescent probe bis-ANS, as it combines with a-lactalbumin. They concluded-not only from these experiments, but also on thermodynamic grounds, as well as from results of adsorption experiments of the apo and bound forms to phenyl-Sepharosethat both Na(1) and Ca(I1) induce a conformational change in alactalbumin, in which hydrophobic regions are removed from the solvent to form a less hydrophobic protein. (For earlier use of bis-ANS in the study of a-lactalbumin, see Section V1,D.) Other uses of fluorescence in examining conformational changes that occur on complexing with metal ions were discussed in Section VI. The results of Kronman et al. (1981), Murakami et al. (1982), Murakami and Berliner (1983), Kronman and Bratcher (1984), and Ostrovsky et al. (1988) have been particularly significant. 4 . Raman Spectroscofi

The Raman spectroscopy of lysozyme dates back more than 50 years, when Edsall(l938) proposed such a study. However, the first spectrum was not obtained by Garfinkel and Edsall(l958) until 20 years later. This was probably the first published Raman spectrum of a biopolymer. More recent studies (e.g., M. C. Chen et al., 1974) resulted from the impetus of the work by R. C. Lord (see Lord and Yu, 1970). The use of Raman spectroscopy in biochemistry was extensively reviewed by Yu (1977),who included a discussion of lysozyme and a-lactalbumin spectra. Yu (unpublished work quoted by Yu, 1974)found that, as would be expected, there was no sign of conformational change for lysozyme, as revealed by the Raman spectrum, as the pH was decreased from 5.2 to 2.0. In particular, there was virtually no change in the amide I11 backbone region (1220-1300 cm-'). On the other hand, there were changes in amide I11 frequencies and the contour of Raman bands for a-lactalbumin upon the same reduction in pH value. Yu (1974, 1977) also showed, by comparison of the environments of the peptide backbone, as reflected in the amide I11 bands, that there was virtually no difference between the spectra of crystals of lysozyme and a-lactalbumin and their respective solutions. However, the spectra were altered by lyophilization, with respect to main-chain conformations (this is considered again in Section XI). Using the 830/850 cm-' doublet as a measure of the negative state of phenolic oxygen and of the tyrosine environment, Van Dael et al. (1987) studied the effect of low pH on bovine a-lactalbumin Tyr groups. They also examined the stabilizing role of Ca(I1) and Na(1) on the structure and the change in state of Trp residues as the molecule unfolds. While considering vibration spectra, mention should be made of the

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study by Careri et al. (1980), who used IR to study the events that occur during the hydration of lysozyme. [For later developments in this work, see the article by Rupley and Careri, this volume.]

5. Circular Dichroism (CD) and Optical Rotary Dispersion (ORD) CD and ORD (generally discussed by McKenzie, 1970, and Simons, 1981) were often used in early comparisons of lysozyme and a-lactalbumin. Although they suffered from limitations in the equipment then available, it is valuable to consider these results here. In Section XI we discuss some future possible studies in this area. Aune (1968) found that the ORD curve for bovine a-lactalbumin (pH 6.1, 0.1 M KC1) was indistinguishable from that of domestic hen egg white lysozyme (pH 4.5,O.l M KC1) in the region 206-233 nm, but that there were large differences in Cotton effects in the 250- to 233-nm region. Similar results were obtained in CD studies by Kronman (1968). Other workers [including one of the authors (H. McK.)] found differences in the 205- to 240-nm region as well as the 250- to 330-nm region. Before considering the conformational significance of these differences, it is necessary to consider the origin of Cotton effects in the 250- to 3 1O-nm region. Glazer and Simmons (1965) were the first to observe a side-chain Cotton effect near 290 nm in the ORD (and later a positive CD band at 294 nm) of domestic hen egg-white lysozyme, and they attributed it to Trp residues. Following publication of the X-ray crystal structure and amino acid sequence of domestic hen egg-white lysozyme, Teichberg et al. (1970) selectively oxidized Trp-108 and found that the positive ellipticity at 294 nm was abolished, leaving negative contributions in the 265- to 300-nm region from other sources, the most prominent extrema being at 293 and 268 nm. It was considered that the oxidation prevented coupling of transitions in residue 108 with those in Trp residues 63 and 111, which are situated nearby. (The positive band is absent in bovine a-lactalbumin, where residues 59 and 107 are not Trp; these residues are 62 and 111 in hen lysozyme sequence numbering.) Comparative studies of CD of human lysozyme and domestic hen egg-white lysozyme have been made by Halper et al. (197l), and of bovine, camel, guinea pig, and human a-lactalbumin, and domestic hen egg, duck egg, goose egg, and human lysozyme by Cowburn et al. (1972). Although the domestic hen egg-white lysozyme has more Trp residues than human lysozyme, Cowburn et al. found the intensity of the CD band near 293 nm was less in domestic hen egg-white lysozyme than in human lysozyme. They concluded that this is due to cancellation of contributions of opposite sign in the domestic hen protein. The 293-nm band in the

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a-lactalbumin is small and is superimposed on larger negative bands from other sources. Cowburn et al. do not agree with the interpretation (based only on the CD of domestic hen lysozyme) of Teichberg et al. (1970) for the 293-nm perturbation and conclude that it involves, at least for human lysozyme, mainly Trp-64 (residue 63 on domestic hen lysozyme numbering). They also consider that the differences in this region between the hen lysozymes and other proteins studied may arise from the adjacent Trp residues 62 and 63 in the hen protein. All of the a-lactalbumins and lysozymes studied, except the egg lysozymes, have negative bands near 270 nm. It is not possible to allocate these bands unequivocally, but they appear to involve Tyr perturbation. Differences in chirality of the disulfide bridges in a-lactalbumin and lysozyme may also cause appreciable contribution to bands in the 240- to 270-nm region. T h e curves in this region cannot be classified simply as typical of mammalian lysozymes, egg lysozymes, and a-lactalbumins. The spectra for bovine, pig, and kangaroo a-lactalbumins are very similar. On the other hand, the curve for human a-lactalbumin is similar to that for human lysozyme, but different from that of domestic hen lysozyme. T h e curve for echidna lysozyme I resembles that of human lysozyme fairly closely. The side-chain effects of red kangaroo a-lactalbumin and echidna lysozyme I resemble those of human lysozyme (Hopper and McKenzie, 1974). On the basis of CD studies, Robbins and Holmes (1970) reported the following tentative fractions for a-lactalbumin structures: 0.25-0.26 (25-26%) of the chain length as a helix, 0.14-0.15 (14- 15%)as /3 structure, and 0.60 (60%)as unordered structure. These values are somewhat different from those reported for lysozyme by Greenfield and Fasman (1969) and by Y.-H. Chen et al. (1974). For the former the fractions were 0.29 (29%) helix, 0.1 1 (11%)p structure, and the remainder unordered. For the latter the values for helix and p structure were 37% and 11%, respectively. X-Ray analysis of lysozyme (Blake et al., 1967a) yielded 28-42% helix and 10% /3 structure. (This range of values for helix indicates regular a helix on the low side, and 3,, as well as distorted helices on the high side, included together with the a helix.) Robbins and Holmes ( 1970) stressed the interpretation that their comparisons meant “similar” conformations and that they had, in effect, corroborated the work by Kronman (1968). On the other hand, Bare1 et al. (1972), using ORD and CD, found slightly more helix for lysozyme than for alactalbumin. More recently, Nitta et al. (1984) reported 33% and 1776, respectively, for a helix and p structure of bovine a-lactalbumin. Thus, while values

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for helix in a-lactalbumin have been within the range found by X-ray analysis of lysozyme, there have been considerable differences in reported values for p structure. Rawitch (1972) found a difference in rotational diffusion of these two proteins and concluded that the effective molecular volume of a-lactalbumin is greater than that of domestic hen lysozyme. Takesada et al. (1973) used CD in a study of the unfolded proteins and concluded that the two differ, with much structure remaining for a-lactalbumin that was not present for lysozyme (see also Section IX,E). CD has been used also in the search for conformational intermediates, in a comparative study of the oxidative refolding of lysozyme and a-lactalbumin (Kuwajima et al., 1985). This topic is dealt with in Section IX,E.

B . Small-Angle X-Ray Scattering Small-angle X-ray scattering measurements yield information relating to protein shape and hydration, as well as radius of gyration, but results based on such studies for a-lactalbumin and lysozyme have been interpreted differently by two groups of workers. Krigbaum and Kugler (1970) concluded that there are appreciable differences in the shape and hydration of the two proteins, but their conclusion has been disputed by Pessen et al. (197 1) and by Achter and Swan (1971). The latter group recalculated the results of Krigbaum and Kugler, making allowance for the effect of a small amount of protein polymerization, mainly dimerization, under their conditions of measurement. When this was done, Achter and Swan concluded that the small-angle X-ray results show the conformations of bovine a-lactalbumin and domestic hen lysozyme to be essentially the same. Although Pessen et al. (197 1) used newly developed apparatus to improve the small-angle X-ray scattering method, they could not reproduce the differences found by Krigbaum and Kugler (1970) and concluded that, except for a small difference in the extent of hydration, the two proteins have essentially identical macromolecular parameters. C . Electron Spin Resonance and Nuclear Magnetic Resonance

ESR and NMR, as is true of fluorescence spectroscopy, have often been used in the study of conformational changes in a-lactalbumin that occur upon complexing with metal ions (Section VI). These methods, however, have been used extensively in studies on lysozyme, as well as on a-lactalbumin. Thus, McDonald and Phillips (1969) used proton

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NMR to study lysozyme (Section VI). They concluded that Co(II), bound at a single site in lysozyme, could perturb resonance positions of protons throughout most of the protein molecule. Cowburn et al. (1970), with use of high-frequency proton NMR as above, as well as ORD, CD, and IR spectroscopy, undertook comparative studies between a-lactalbumin and lysozyme and found results consistent, with close conformational similarity. The observed differences between the spectra of these proteins were considered to be almost entirely explained by sequence differences. Other ESR and NMR studies have also been dealt with in Section VI. Gallo et al. (1971) and Teichberg et al. (1974) used ESR to explore the binding of metals to lysozyme. In addition, Berliner et al. (1983) used NMR as well as ESR to study the Mn(I1) binding site of a-lactalbumin. More recently, Musci et al. (1987)used ESR and NMR to determine certain intramolecular distances between spin-labeled Met-90 and the metal binding site, as well as certain resolvable protons. Bradbury and Norton (1975),on the basis of the model building studies by Browne et al. (1969) and by Warme et al. (1974),were able to make assignments of specific resonances in the proton NMR spectrum of bovine a-lactalbumin to the three His residues. These resonances, after reaction of the protein with iodoacetate under conditions that were nearly specific for the His residues, disappeared from the “native” frequency positions, but did so in a differential manner, consistent with differences in the degrees of exposure to the solvent. This was predicted by the model building studies, particularly those by Browne et al., which were subsequently confirmed by X-ray analysis. Thus, His-68 is the most exposed; His-32, being involved in a helical region (according to Browne et ul., but not to Warme et ul.), is less exposed, while His-107 is the least exposed. NMR has found more recent use in comparative studies of lysozyme and a-lactalbumin. Poulsen et al. (1980), using the nuclear Overhauser effect, were the first to demonstrate the existence of the “hydrophobic box’’ region, in solution, for lysozyme, first noted in the crystalline state by Blake et al. (1967a). Koga and Berliner (1985) applied the nuclear Overhauser effect to the study of a-lactalbumin. They too found a hydrophobic box, similar to that of lysozyme. The study was conducted in both the presence and absence of Ca(II), and there were only “subtle” differences upon removal of the cation. Berliner and Kaptein (1981) used another NMR method, induced dynamic nuclear polarization, to investigate the solvent accessibility of Tyr, Trp, and His residues in five species of a-lactalbumin. This method measures the access of the photo-excited flavin dye to the surface-exposed

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residues. Many differences were noted among the various residues of the protein examined, but Tyr-50 was the only one consistently unexposed. This finding appears to be consistent with the results of Acharya et al. (1989), in which Tyr-50 was placed inside a pocket of residues containing Ile-41, Thr-48, Asn-57, Ser-63, Leu-81, and S-S bond 61-77. The binding of monosaccharide inhibitors to hen egg-white lysozyme was studied with 270-MHz NMR by Perkins et al. (1981) (see Section IV). There is increasing use of NMR in the study of protein conformation (Gronenborn and Clore, 1990). Its application to the study of conformational changes upon modification of a-lactalbumin is considered in Section IX,E.

D . Association and Aggregation The effects of pH on association and aggregation of both lysozyme and a-lactalbumin have long been known. Sophianopoulos and Van Holde (1961) found lysozyme to be monodisperse at pH 5.4 and 20°C; but at more alkaline pH values the protein dimerizes. Sophianopoulus and Van Holde (1964) and Deonier and Williams (1970) reported detailed studies of the dimerization, which is rapid and reversible at Z > 0.0 1, carried out with sedimentation equilibrium and viscosity studies. The results are consistent with the hypothesis that dimerization is favored by the loss of a single proton from the monomer. The viscosity results of Sophianopoulos and Van Holde do not suggest significant changes in tertiary structure of the protein during the dimerization. However, the near-UV CD exhibits concentration dependence on association of lysozyme (Holladay and Sophianopoulos, 1972), and in the region of 256-293 nm reflects changes in the Trp residue. The rapid reversible association and slow aggregation of bovine alactalbumin at pH <4.0 have been studied in detail by Kronman and coworkers (e.g., Kronman and Andreotti, 1964; Kronman et al., 1964). They found that the protein was largely insoluble in the region of pH 4.2-5.2 and soluble at higher pH values. It was very little associated at pH 6.0 and even less at pH 8.5, the conformation being constant in the pH range 6.0-9.5. Kronman et al. (1967) showed that bovine a-lactalbumin undergoes some expansion, without aggregation, above pH 9.5. They concluded that these phenomena are related to a “denaturation-like” process. Robbins et al. (1965) sought to test the hypothesis that the acid aggregation of a-lactalbumin was due to hydrophobic interactions. Their approach was to insert additional nonpolar residues by amidination, and then examine the acid behavior of the product. They found the modi-

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fied protein to be much more susceptible to aggregation than was the native protein. Their results lend support to the hypothesis that hydrophobic interactions play an important role in the aggregation of alactalbumin. When all of the above work is considered, it is obvious that there are marked differences in association and aggregation behavior between lysozyme and a-lactalbumin, but the structural basis for these differences is poorly understood.

E . Denaturation and Renaturation The denaturation of proteins is discussed elsewhere by Tanford (1970); McKenzie and Ralston (1971); and McKenzie (1991). We concern ourselves here with some aspects of a-lactalbumin and lysozyme. Before discussing denaturation it is desirable to recapitulate work on transitions in a-lactalbumin and the variation in nomenclature that has been used. Following their early studies on the effect of pH on bovine a-lactalbumin, Kronman et al. (1972a,b) made more extensive studies of the effect of pH and temperature on both bovine and caprine a-lactalbumin. For the latter they classified the transitions as follows: Transition I: A pH-dependent transition in the pH range 4-8 involves changes in the environment of Trp residues, but does not involve any change in shape, nor is there any extensive conformational shift. Transition 11: In the pH range 2-4 transition I1 involves some conformational shift and expansion of the molecule. This was called the N + U conversion and is now the N + A conversion, as indicated in Section VI. At 25°C there is no increase in Trp group exposure, although there are changes in the environments of these groups. The results at 3°C are ambiguous. Transition IIA: There is an expansion of the molecule above pH 9.5. It is unresolved as to what extent this change is similar to that involved in transition 11. Regardless of whether the protein is subjected to pH values <4.0 or >9.0, is heated above 50”C, exposed to low concentrations of guanidine hydrochloride, or subjected to Ca(I1) removal from the N form, all usually appear to involve dissociation of Ca(I1) in some way. The original use of the term “U” by Kronman et al. (1972a,b) for altered forms of the protein differs from that of other workers more recently, whereby the term has been taken to designate completely un-

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folded protein (see, e.g., Baum et al., 1989, for the unfolded state in 9 M urea at pH 2.0). T h e term “U” with numerical subscripts (e.g., U 1 , U p , and U,) has also been used in various discussions on denaturation of various intermediates in unfolding. For transition I1 and similar types of transitional change, the term “A state” is now used by most workers and appears to have been used first by Kuwajima (1977). Denaturation of a-lactalbumin appears to be a three-state process (Kuwajima et al., 1976, 1985; Kuwajima, 1977). The initial folding of a-lactalbumin to the intermediate stage, assumed to be essentially the same entity as that occurring on denaturation, is dependent on local interactions, followed by hydrophobic interactions, and long-range specific interactions (S-S bond formation and electrostatic attractions). S-S bonds are not important for stabilizing the intermediate. Kuwajima and colleagues point out that their folding model is not inconsistent with the thermodynamic hypothesis of Anfinsen and Scheraga (1975), since the native conformation may yet prove to have the lowest Gibbs free energy; nevertheless, it may be reached through kinetically controlled intermediates. However, according to Rao and Brew (1989), an additional requirement is the presence of Ca(I1) for the refolding of a-lactalbumin, and this cation is essential for the correct pairing of half cystine residues to form disulfide bonds, as well as for the development of native conformation. Dolgikh et al. (1985) suggested a compact intermediate with a slowly fluctuating tertiary structure for a-lactalbumin. Ptitsyn et al. (1983) proposed a model for a-lactalbumin whereby the folding intermediate was seen as a compact globule with fluctuations in its three-dimensional structure (see also Shakhnovich and Finkelstein, 1982). More recently, Gil’manshin et al. (1988) found an early intermediate in the folding of a-lactalbumin that forms in sec after denaturation with 8 M urea. This time is two orders of magnitude smaller than that found by Kuwajima et al. (1985). T h e latter authors maintained that their folding intermediate had essentially the same secondary structure as their denaturation intermediate (the A state). Kuwajima (1989) reviews findings on the “molten globule state” and suggests that it may be a common state among proteins. Baum et al. (1989), through an NMR study of the A state of alactalbumin, produced a more detailed structural picture than has been seen previously. Methods were used that exploit the well-resolved spectrum of the native state to examine the A state indirectly. Some resonances in the latter were considerably shifted from their native positions and were identified through magnetization transfer with the native state.

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Of particular interest was the apparently unchanged helical segment 89-96 in the A state, as indicated by a number of amides that were protected from solvent exchange. It was suggested that the stability of this segment may be brought about by a variety of side-chain interactions. From their results Baum et al. were able to propose a model for the A state, in which much conformational freedom is exhibited, but in which specific elements of the native state are preserved. The denaturation of lysozyme, on the other hand, is a two-state process, according to Tanford (1970). In addition, there is high cooperativity between S-S bonds and conformational structure for this protein, compatible with the hypothesis that the formation of native S-S bonds may occur as an essential prerequisite early in the folding process, prior to the onset of native conformational structure (White, 1982; White and Wright, 1984). The work by Galat et al. (1981), in fact, indicates that, after at least three of the four S-S bonds reform, all measured elements of conformation appear simultaneously. On the other hand, Kato et al. (1981) found spectral evidence for a “rapidly” formed structural intermediate in the refolding of hen egg white lysozyme. It was concluded that the unfolded species assumed its new transient conformation in the mixing process of the pH-jump measurements and that the transformation was complete within the mixing time. Unfolding had been achieved by either 4 M guanidinium chloride or 40% acetic acid. Further evidence for the essential nature of S-S bond formation in the development of native conformational structure, for human lysozyme, comes much more recently from Taniyama et al. (1988). They produced the first evidence that formation of a specific disulfide bond (Cys-56-Cys-28) is a prerequisite for the correct folding of this protein in vivo, expressed in Saccharomyces cerevisiae. Notwithstanding the suggestions of Kuwajima and co-workers for the renaturation of a-lactalbumin (above),it is difficult to see how the renaturation of lysozyme in particular could be governed primarily by thermodynamic influences, in view of the reported requirements for native S-S bonds (for further discussion see White, 1982; White and Wright, 1984). Additional evidence of difference in the refolding process is that the time taken for reoxidation of a-lactalbumin is longer than 25 hr, whereas that for lysozyme is only 90 min (Tamburro et al., 1972). Iyer and Klee (1973) found differences in the rates of S-S bond reduction in these proteins. Reduced a-lactalbumin retained hydrodynamic and optical properties characteristic of folded globular proteins, although its conformation was clearly distinguishable from that of the native protein. Evidence of sameness in the denaturation of a-lactalbumin and lyso-

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zyme comes from Sharma and Bigelow (1974), according to whom these proteins behave similarly on denaturation, and therefore would be expected to have similar backbone conformations (cf. Maes et al., 1969; Bradbury and King, 1971). An indication of sameness on reoxidation comes from Chiaranda et al. (1972), who attempted to reoxidize a reduced mixture of two CNBr fragments of or-lactalbumin; the CD spectra that appeared did not suggest that the native conformation could have been reformed. This observation was interpreted to mean that S-S bonds form early in the renaturation of a-lactalbumin, consistent with the previously mentioned work on lysozyme (Galat et al., 1981; White, 1982; White and Wright, 1984; Taniyama et al., 1988). Thus, the two proteins appear to be similar in this respect. In this picture the differences appear at least as striking as the similarities, in comparing both denaturative and renaturative properties of a-lactalbumin and lysozyme. Such differences are compatible with differences in the conformations of these proteins in the native state, in aqueous solution. T h e behaviors of apo- and Ca(I1)-bound forms of a-lactalbumin differ markedly upon denaturation with guanidine hydrochloride, as shown by Ikeguchi et al. (1986). Thus, at low Ca(I1) ion concentration alactalbumin unfolds to produce a stable intermediate, while at high Ca(I1) concentration the protein unfolds in a manner similar to that of lysozyme. Further studies on the conformers of a-lactalbumin were reported by Hanssens et al. (1984). The thermal transition curve differs between the Ca(I1) and apo forms of a-lactalbumin, although these forms have similar fluorescence characteristics. F. Chemical Reactivities Reactivity differences between a-lactalbumin and lysozyme are given by Lin (1970), where the carboxyls of these proteins are concerned. These differences, however, were considered to be compatible with the model of Warme et al. (1974). Atassi et al. (1970) found S-S bonds in a-lactalbumin to be more susceptible to reduction. Barman and Bagshaw (1972) found six peptide bonds in a-lactalbumin to be digestible by trypsin, but not so in lysozyme, which is therefore relatively resistant to this enzyme. Further, these workers concluded that Trp-26 (also Trp-104 and Trp-108) is not buried-but it is, according to the model of Browne et al. (1969) and the crystal structure. Takesada et al. (1973) made several observations: (1) the number of

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slowly exchanging peptide hydrogens was 35 in a-lactalbumin, but 44 in lysozyme; (2) kinetic profiles of the exchange reaction were different for the two proteins; (3) the midpoints of their thermal transitions were different; (4) effects of pH on the exchange kinetics differed; ( 5 ) a CD study indicated that the unfolded form of a-lactalbumin has an “appreciable amount” of folded structure. A similar conclusion was reached by White (1976) for lysozyme, after full reduction of S-S bonds in 8 M urea, followed by redissolving in dilute buffer. More recently, attention has focused on a specific residue in a-lactalbumin as being essential for its activity in combining with galactosyltransferase to form the lactose synthase system. This is His-32, which is consistently present for all species, as would be necessary for an essential residue. Schindler et al. (1976) specifically modified this residue with diethyl pyrocarbonate, to give the ethoxyformyl derivative, with inactivation of the protein. This His lies within the cleft region of a-lactalbumin and in this position might substitute for Glu-35, which is present in the lysozymes (Fig. lo), thus possibly accounting for the trace of cell lytic activity reported by McKenzie and White (1987). This activity is considered in more detail in Section X. Pfeil (1981) concluded that a-lactalbumin is less stable than lysozyme, with a lower thermal transition temperature, lower denaturational enthalpy, lower heat capacity change, and lower Gibbs free-energy change. Although these generally greater reactivities for a-lactalbumin may suggest a “looser” conformation for this protein, such may not be the case, according to Barman (1970),who suggested two conformations for a-lactalbumin in equilibrium in the native state: one, a relatively tight globular conformation; the other, a more diffuse conformation, which likely would be more accessible to various reagents and to denaturation in general. Thus, when the latter undergoes change, the equilibrium shifts in the direction of the more open conformation.

G. Immunochemical Properties Tanahashi et al. (1968) compared the immunological properties of bovine, water buffalo, ovine, caprine, porcine, guinea pig, and human a-lactalbumins by the method of Oudin. They found that the nonruminant a-lactalbumins do not react with antisera to the bovine protein. This is in accordance with our experience (K. Bell and H. A. McKenzie) using the Ouchterlony and immunoelectrophoretic methods. However, Sakar et al. (1971) found that while bovine, water buffalo, and caprine a-lactalbumins exhibit extensive cross-reaction in the Ouchterlony test, these ruminant a-lactalbumins may be differentiated quanti-

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tatively in the microcomplement fixation method. Atassi et al. (1970) found that bovine a-lactalbumin and domestic hen egg-white lysozyme do not show cross-reaction in quantitative precipitin analyses. However, weak cross-reaction in hemagglutination determinations was found by Strosberg et al. (1970). On the other hand, Faure and JolKs (1970) found no cross-reaction in similar studies for bovine a-lactalbumin, human milk lysozyme, and various egg lysozymes against anti-a-lactalbumin sera. The only antigenic similarities observed in “assays” against antidomestic hen lysozyme sera were among the several egg lysozymes. Arnheim et al. (1971) found that there was no cross-reaction between the native forms of human and domestic hen egg-white lysozymes, but there was cross-reaction between the reduced carboxymethylated proteins. They discussed several reasons for this and pointed out that antisera produced against unfolded proteins can be useful in evolutionary studies, but they cautioned against pitfalls in this approach. One application is that of Arnon and Maron (1971), who found cross-reactivity between the reduced carboxymethylated forms of a-lactalbumin and lysozyme. They regarded this observation as corroborating evidence that the two proteins evolved from a common evolutionary precursor. Light has been shed by the work of Gavilanes et al. (1984) on the question of why the native forms do not cross-react. Thus, secondary structures of lysozyme and a-lactalbumin, predicted on the basis of the method of Chou and Fasman (1974), are sufficiently different in the region of the antigenic loop (residues 60-83 in hen egg-white lysozyme) that it is not expected that the native forms of these proteins would cross-react. It appears to be generally accepted that, at least for lysozyme, most of the antigenic determinants are assembled topographic determinants. This concept is supported by the observation that little or no crossreactivity occurs between native and denatured lysozymes (Thompson et al., 1972). The antigenic determinants of the native lysozyme molecule appear to include most, if not all, of the surface residues, as evidenced by numerous studies (reviewed critically by Benjamin et al., 1984). This point of view contrasts with that of Atassi and Lee (1978), who claimed a limited antigenicity, based on a study of “surface-simulated peptides.” The sites delineated by the latter workers do not include Arg-68 (Fainanu et al., 19?4), the “loop” region in general, or any of several other segmental regions previously demonstrated to function in this capacity, such as residues 1-20 and 123-129 (see Benjamin et al., 1984). Hopp and Woods (1982) attempted to locate the antigenic sites of a-lactalbumin and found that this activity can be attributed to several peptic fragments and to single Arg and Met residues. They concluded

274

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

that although hen egg-white lysozyme does not cross-react with the a-lactalbumins studied, there are similar distributions of antigenic determinants on the surfaces of these a-lactalbumins and lysozyme. Smith-Gill et al. (1982), with the aid of a monoclonal antibody (HyHEL-5) prepared to domestic hen egg-white lysozyme, were able to identify an antigenic site (i.e., epitope) that was shared by the lysozymes of seven species of galliform birds. Two of these, bobwhite quail and chachalaca, shared only partial identity with epitope defined by this antibody. Duck lysozyme did not react with the antibody. Assuming a lack of long-range conformational changes, Arg-68 was identified as a determining residue. Together with Arg-45, to which it is hydrogen bonded, Arg-68 formed a basic cluster which may be a subsite of the epitope. Further evidence indicated that the epitope extended into the cleft region between Arg-45 and Arg-114 (Fig. 11). Using both the X-ray structure of the Fab :lysozyme complex (Sheriff et al., 1987, 1988; Amit et al., 1986) and the site-specific mutagenesis of

b

LOOP

HyHEL-5 determinant Catalytic Cleft HyHEL-12 HvHEL-7

525.5E4 Hyb.Cl

I

I

HyHEL-9 525.3c7

325.3D1 (residue 121) HyHEL-11

FIG. 11. (a) The amino acid sequence of domestic hen egg-white lysozyme, showing eight peptides that have been shown to be antigenic against anti-domestic hen egg-white lysozyme. N-C peptide is indicated by solid outline; LH,, by a dashed outline; Plb, by stippling; a continuous region (residues 34-54) within Plb, shown as black box; peptide 8, by a heavy black outline; the loop, by black with white lettering; loop 11, by a stippled box; LIII, by a dotted outline. (b) “Space-filling’’model of domestic hen egg-white lysozyme (computer generated). The loop and N-C peptide are dark gray, with residues 1-3 black. Specific residues recognized by monoclonal antibodies are colored or outlined in white: a hypothetical unit determinant for antibody HyHEL-5 is outlined in dotted black. [Reproduced with permission from Benjamin et al. (1984), who give details of the antibodies in their Fig. 2.1

LYSOZYME AND

a-LACTALBUMIN

275

a hen egg-white lysozyme cDNA gene (expressed in yeast), an epitope containing Arg residues 45 and 68 was found by Lavoie et al. (1989b), confirming the original prediction from epitope mapping of evolutionary variants of egg-white lysozymes (Smith-Gill et al., 1982). Site-specific mutagenesis, in particular, is of continuing value to these workers in their exploration of the interaction of monoclonal antibodies with lysozyme. Lavoie et al. (1989a) also compared the crystal structures of two monoclonal antibodies with serological data, and for one of these (their HyHEL-5, complexed with lysozyme) the results agree. However, for their HyHEL-10, complexed with lysozyme, similar comparison does not result in agreement. For the former, however, Arg residues 45 and 68 are implicated as epitopes. Thus, this conclusion is essentially in accordance with that of Lavoie.

H . Conclusions The studies summarized in this section have proved particularly valuable, because they were conducted in (mostly aqueous) solution, enabling comparison with X-ray or neutron diffraction studies, for which crystalline material must be used. Although the conformation of a crystalline protein can be essentially the same as that in aqueous solution, it is not necessarily correct to make this assumption, since there may be large conformational changes in such solution, contingent on the variables of protein concentration, pH, ionic strength, and temperature. In fact, there are many reported differences between a-lactalbumin and lysozyme in solution, in comparison with the similarity between these proteins indicated by high-resolution X-ray analysis of their crystals. On the other hand, UV absorption and fluorescence methods have been involved in relatively few comparative studies between a-lactalbumin and lysozyme. More useful to this end have been CD and ORD, both of which reflect changes in various short-range interactions. Also noteworthy have been the many studies with ESR and NMR. These, in addition, have found considerable use in the study of complexing between a-lactalbumin and metal ions. They will presumably be of value in the future for the study of such reactions for those few lysozymes that coordinate to metal ions. Other methods, including Raman spectroscopy and small-angle X-ray scattering, have yielded results generally supporting similarity between the conformations of a-lactalbumin and lysozyme. There may be two reasons for the reported evidence of dzfferences in conformation and reactivity in aqueous solution: (1) substantial differ-

276

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

ences in conformational structures of these proteins may indeed exist in solution; and (2) the suggestion of Barman (1970; see also Kuwajima, 1977; Dolgikh et al., 1981) may prove to be correct (i.e., a “tightly” folded conformation may exist in equilibrium with a form of a-lactalbumin that is more reactive). Such a phenomenon would explain why a-lactalbumin appears to be more accessible to various reagents than is lysozyme, and why it is more easily denatured, despite indications of a close conformational similarity. Thus, the reactive form of the protein is altered, whereupon equilibrium between the two forms is continually shifted to produce more of the latter form, which then continues to react. If this idea proves to be correct, the question arises as to what specific structural features of the “tightly” folded form of a-lactalbumin (compared with lysozyme) permit transformation to the more reactive form.

X. EVOLUTIONARY ORIGINS OF LYSOZYME AND a-LACTALBUMIN A . Introduction: Molecular Clocks and the Fossil Record As is obvious from a number of recent attempts to confront aspects of evolutionary history and evolutionary mechanisms, there is at present a greater openness to discussion than at any period since the formulation of the “Modern Synthesis”. This is largely the result of the revolution in molecular biology which has removed many of the restrictions on discussion of evolutionary mechanisms accepted since the development of that synthesis. But it has also followed on a period of reinterpretation of the fossil record as a source of new historical data, and as a testing ground for mechanisms. The claim of the palaeontologists to exclusive rights to the interpretation of the sequence and dating of evolutionary events has been under siege by advocates of cladistic taxonomy and, more effectively, from proponents of molecular clocks. Finally, the much publicised attack on the logical status of many of the explanatory concepts of Neo-Darwinism has pushed many workers into re-examining possible explanations for evolutionary phenomena that would have had few advocates five years ago. In the resulting turmoil, and it is indeed a turmoil, the once dominant fields of population genetics and development biology have provided a stabilising influence, rightly pointing to experimental data on the factors influencing evolutionary processes to counter some of the less well grounded and more speculative views.

From the preface by Campbell and Day (1987) to a ‘Rates of Evolution’ symposium

1 . Molecular Clocks The above comments reflect the trepidation that we feel as chemists, who do not profess to be either paleontologists or evolutionary biologists, in discussing this wide-ranging problem. It is evident in what has already been presented in this article that a-lactalbumin and lysozyme have important evolutionary relationships that involve divergence and/

LYSOZYME AND (Y-LACTALBUMIN

277

or convergence. Before discussing specifically their relationships, we must consider some important general background aspects. Even 30 years ago paleontology was virtually the only source of information about the periods when common ancestors lived. Indeed, the mammalian fossil record was not particularly good. Early writers on comparative biochemistry, such as Baldwin and Florkin, were limited in their sources. Perhaps the first person to perceive the unique value of the molecular evolution of proteins and nucleic acids was Anfinsen (1959), who wrote of this subject in his important book. However, it appears to have been Zuckerkandl and Pauling who, in 1960-1965, introduced the concept of the molecular clock (for a historical review which is part of a group of important critical papers in an issue of the Journal of Molecular Evolution, see Zuckerkandl, 1987). Zuckerkandl and Pauling (1962) and Margoliash (1963), by making sequence comparisons of hemoglobins and cytochromes c, respectively, concluded that the number of point mutations needed to account for the differences in amino acid sequences is correlated linearly with estimates from paleontology of the time elapsed since the species compared had a common ancestor. In discussing the molecular evolutionary clock, it is important to stress that it does not have the accuracy of the National Bureau of Standards standard of time, let alone that of radioactive decay. Furthermore, it is not absolute, but must be calibrated against fossil records. Many workers have reported that the amino acid sequences of proteins usually diverge at fairly constant rates. Often, the point mutations have no effect on protein function. Wilson et al. (1977), in their important review, indicated that each functional class of proteins tends to have its own specific rate of change. They used the term “unit evolutionary period,” which is the time, in units of lo6 years, required for the accumulation of a 1% difference in amino acid sequence. Concomitant with such studies, Kimura developed his neutral theory of molecular evolution. He asserted that the great majority of evolutionary changes at the molecular level, as seen in comparative studies of protein and DNA sequences, are the result not of Darwinian selection, but of random drift of selectively neutral (or nearly neutral) mutants (for reviews see Kimura, 1983, 1987). While such views were an anathema to some biologists and geologists, perhaps even greater concern was expressed over the views of Wilson and others on the time dependence evident in quantitative immunological comparisons of mammalian albumins and also of lysozymes (Wilson et al., 1977). One important factor in this lack of acceptance was the faulty view of the antigenic structure of proteins existing in the early 197Os,when much of this work was done (as we have already seen for lysozyme and a-lactalbumin in Section

278

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

IX,G). However, with more recent ideas on antigenic structure, as reviewed by Benjamin et al. (1984), apparently there is evidence that a good correlation may exist between immunological distance and sequence difference for some globular proteins. Hence, valuable information can be obtained about rates of change for many mammalian groups, as well as some plants, bacteria, etc. (see also the general review by Wilson et al., 1987). Despite the considerable body of evidence that Kimura accumulated in support of his neutral theory, he did find it necessary to reconsider some departures from clockwise progression of molecular evolution and to suggest future experimental programs in an attempt to settle certain issues (Kimura, 1987). In addition to comparison of amino acid and nucleotide sequences of a-lactalbumin, and their rates of change as molecular clocks, a considerable amount of comparative information has accumulated on the three-dimensional structure of these proteins; their physical properties in solution; effects of amino acid substitutions, in both genetic and cloned variants; and their functions and immunological properties. In assessing this information it is important not to lose sight of the known paleontological information on the origin and evolution of mammals. 2. Mammalian Evolution and Paleontology

The evolution of mammals has not been without controversy: Over 20 years ago Crompton (1968) entitled an important article, “The Enigma of the Evolution of Mammals.” Nevertheless, the majority of paleontologists would accept the phylogenetic relationships of the major groups of amniote tetrapods-reptiles, birds, and mammals-summarized diagrammatically in Fig. 12a. Clemens (1989) has maintained that, although one recent comparative study resulted in Gardiner’s (1982) proposal for a closer relationship between birds and mammals (see Fig. 12b), the weight of the evidence, particularly that from the fossil record, supports the interpretation given in Clemens’ article (see also other references in Clemens, 1989; for general discussions see Benton, 1984, 1990; Carroll, 1988; Gauthier et al., 1988). A critical feature of Clemens’ view (summarized in Fig. 12c) is the basal dichotomy between the group including modern reptiles and birds on the one hand and that including modern mammals on the other, and now appearing to date back some 300 million years. The Monotremata are seen as part of the other major branch of the diagram. The discovery of Steropodon in Australia extended the record of Monotremata back to -100 million years ago (Archer et al., 1985). A theme underlying many mammalian classification studies has been the desire to determine better the relationships of the modern monotremes, marsupials, and eutherians. The origin of the monotremes

I &' Ha

othermia

E"+

A

T

iota

I Tetrapoda

(b)

Mammalia

Mammal like "reptiles" A (ca. 300) C

FIG. 12. Some evolutionary relationships based on paleontological evidence. (a) and (b) Two views of the evolution of tetrapods (amphibians, reptiles, birds, and mammals) (a) shows a 'conventionay view (see, e.g., Benton, 1984, 1990); (b) shows the view of Gardiner (1982). Common names are shown on (a), systematic names in (b). (c) Clemens' (1989) view of the phylogenetic relationships of major groups of amniote vertebrates (birds, reptiles, mammals), especially showing monotremes, marsupials, and eutherian mammals. Divergence at A probably occurred about 300 MY ago (late Carboniferous). Divergence at B being about 135 MY ago (late Jurassic). Date of divergence at C is not known (although some consider 200 MY ago, late Triassic). D: Steropodon, dated about 100 MY ago (early Cretaceous). E: 90- 100 MY ago.

280

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

and their relationships to Marsupialia and Eutheria, in particular, were discussed by Griffiths (1978, 1989). From these and other studies (see, e.g., Kemp, 1983; Clemens, 1989) the therian mammals, including Marsupialia and Eutheria, still appear to be regarded reasonably as a monophyletic group. However, the former differentiation placing monotremes in a separate group may no longer be valid. [For general background information on marsupials and monotremes, see the articles by Dawson (1977) and by Griffiths (1988), respectively.] B . Evolution of Lysozyme and a-Lactalbumin: Divergence andlor Convergence? Even when Brew and Campbell (1967) suggested, on the basis of similarities in amino acid composition, that a-lactalbumin and lysozyme have diverged from a common precursor, proposals for the evolutionary development of homologous proteins were not new. Examples then known included pancreatic zymogens (Hartley et al., 1965), hemoglobin and myoglobin (Ingram, 1963), and immunoglobulins (Hill et al., 1966). However, none of these has the wide appeal of a-lactalbumin-lysozyme divergence because of the development of their contrasting biological functions. Subsequent comparisons of a wide variety of a-lactalbumins and ctype lysozymes revealed 35-40% identity in amino acid sequences; similarity, but not identity, in three-dimensional structures; high conservation of disulfide bridges; and similarity in many other properties. Such studies have resulted in the general acceptance of divergence of a-lactalbumin and c-type lysozymes, rather than their convergence. In the latter type of evolution, two unrelated genes may have evolved until arriving at a stage at which homology exists in the corresponding proteins, with the resulting sequences and conformations sufficiently adapted to perform their respective biological functions. The argument against this convergent process appears to be strong, and it has been further strengthened by the study of introns and exons. 1 . Zntrons and Exons In contrast to the genes of prokaryotes, for which the coding sequences are continuous, those of eukaryotes are present in blocks (exons) separated by intervening noncoding sequences (introns). Gilbert (1978) who introduced these terms, suggested that exonhntron structure could provide a mechanism for increasing the rate of evolution. It was pointed out that if, for example, exons corresponded to units of protein function, recombination within introns could reassort protein functions to

28 1

LYSOZYME AND a-LACTALBUMIN

produce new proteins from parts of existing ones. Furthermore, Blake suggested that if exons also corresponded to integrally folded substructural domains, there would be a higher probability that this type of recombination would result in a folded viable protein molecule (for a review see Blake, 1983). Comparisons of lysozyme and a-lactalbumin have been made at the transcriptional level. Thus, cDNA clones and nucleotide sequences of rat (Dandekar and Qasba, 1981; Qasba and Safaya, 1984), guinea pig (Craig et al., 1981; Hall et al., 1982), human (Hall et al., 1987), caprine (Kumagai et al., 1987), ovine (Gaye et al., 1987), and bovine (Vilotte et al., 1987; Hurley and Schuler, 1987) a-lactalbumins are now known, together with much of their flanking sequences. It can be seen from Table X that the genes of the four a-lactalbumins shown consist of four exons, separated by three introns. The introns occur at identical positions in all four species. Although the sizes of the exons are highly conserved, those of the introns are not. As would be anticipated, DNA sequence comparisons reveal much homology within the exons, but less conservation in the introns. If domestic hen egg-white lysozyme (Jung et al., 1980), human lysozyme (Peters et al., 1989), and bovine stomach (Irwin et al., 1989) are included in the comparison, introns occur at positions in the genes for the two lysozymes corresponding to those for the four a-lactalbumins (Acharya et al., 1989, comment that such results do not appear to agree TABLE X Comparison of Exon and Intron Lengths" of a-Lactalbumin and Lysozyme Genes

Gene a-Lactalbumins Human Bovine Guinea pig Rat Lysozymes Domestic hen Human lk Bovine st Mouse M

(1) (2) (3)

(4) (5) (6) (7) (8)

159 160 165 165

648 32 1 338 34 1

159 159 159 159

489 473 48 1 429

76 76 76 76

499 504 50 1 1016

333 330 314 328

165 164 >151 166

1270 1563 -2100 -1300

162 165 165 165

1810 1938 1900 -2000

79 79 76 79

79 853 -2700 -450

180 1094 553 854

-

"In base pairs.

bHen,Hen egg white; lk, from leukemic material; st, stomach mucosa. c ( l )Hall et al. (1987), ( 2 ) Vilotte et al. (1987), (3) Laird et al. (1988), (4) Qasba and Safaya (1984), (5) Jung et al. (1980), (6) Peters et al. (1989), (7) Irwin et al. (1989), (8) Cross et al. (1988).

282

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

entirely with the domain theory). Hall and Campbell (1986) point out that the acquisition of introns preceded the gene duplication event that produced a-lactalbumin. The variation of intron lengths among these genes, however, indicates divergence within these regions following gene duplication (see Table X, which also includes results for the mouse M gene). In further examination of these structures, exon 2 has proved to be the most highly conserved and appears to be involved with substrate binding in both proteins, in addition to active site residues in lysozyme. Exon 4 plays a role in galactosyltransferase binding for a-lactalbumin and accordingly shows little similarity to exon 4 for lysozyme, which does not bind with galactosyltransferase. Also, the conserved Asp residues corresponding to exon 3 are consistent with the relatively high affinity of a-lactalbumin for Ca(I1). In their comparison of the nucleotide sequence of the coding region of human pre-a-lactalbumin cDNA with that of domestic hen egg-white prelysozyme cDNA, Hall et al. (1982) found, after maximizing alignment, that there was a greater nucleotide (53%) than amino acid (38%) identity. They also noted that, of the codons that differ by only a single base, more than half represent “silent” substitutions. These observations add further weight to the concept that lysozyme and a-lactalbumin arise from divergent evolution from a common gene, as opposed to convergent evolution from two distinct genes. In their recent study of multiple genes for ruminant lysozymes, Irwin et al. (1989) noted that, throughout evolution, the positions of the introns in the lysozyme gene have remained at identical locations within the lysozyme family. However, differences have been observed in both the lengths of introns and the total gene length. In comparison with a-lactalbumin, lysozyme genes show greater differences, both in the sizes of their introns and in gene length, from each other and from the alactalbumins examined, than was observed within the latter group of proteins. Thus, they suggested the possibility that evolutionary change in size of the genes, from lysozyme to a-lactalbumin, may be associated with the change in gene expression, which may have involved the insertion and deletion of enhancer elements. 2 . Chick-, Goose-, Phage-, and Insect-Type Lysozymes

The homology of a-lactalbumin with lysozyme, the similarity in threedimensional structure and molecular size, etc., are for the well-known c-type (chick) lysozyme. However, there are other forms of lysozyme that catalyze the same reaction as c type. These include insect lysozymes, which are essentially of two types: the c type, (Jolles et al., 1979b; Eng-

LYSOZYME AND (Y-LACTALBUMIN

283

strom et al., 1985) and a type having a high level of sequence identity with phage lysozyme and a high chitinase/muramidase activity ratio (Fernandez-Sousa et al., 1979). Other forms of lysozyme include phage lysozyme (Remington et al., 1978), g-type (goose) lysozyme (Isaacs et al., 1985), and bacterial lysozyme from Streptomyces erythraeus (Harada et al., 1981; see also the review of Jolli3 and Jolli.s, 1984). These non-c-type lysozymes are not our concern here, but some brief comments are pertinent. Three-dimensional structures of the latter three have been determined, and two of them (goose and bacteriophage T4) have been intensively compared with c type by Weaver et al. (1985). Even though their amino acid sequences appear to be unrelated (for early work see Tsugita and Inouye, 1968), Grutter et al. (1983) suggested that the nature of their structural correspondence indicated that c, g, and phage types diverged from a common evolutionary precursor (see also Matthews et al., 1981, and the review by Bajaj and Blundell, 1984). Weaver et al. (1985) noted some similarities in the active site of the three lysozymes, but with the following striking difference. Residue 73 (Glu) in goose corresponds with residue 35 (Glu) in chick and with residue 11 (Glu) in bacteriophage T4. On the other hand, there are two Asp residues at positions 86 and 97 in the goose active site, neither of which corresponds exactly with Asp-52 of chick nor Asp-20 of T4. The implications for potential differences in the mechanism of catalytic action by the three lysozymes were discussed by Johnson el al. (1988) and by Weaver et al. (1985). T h e latter authors discussed the unresolved question as to whether the c-type lysozyme exons correspond to distinct structural and/or functional entities that are conserved during evolution of the three types of lysozyme considered. Phylogenetic analysis of amino acid sequences of c-type egg-white lysozymes from a variety of birds is generally in accord with taxonomic classification. However, there are some differences: For example, the chachalaca is classified normally in the order Galliformes, but its lysozyme differs more in sequence from those of pheasantlike birds than do the c-type lysozymes of ducks (for a discussion of this and other examples, see Jolles and Jolles, 1984). 3’. Stomach Lysozymes: Convergence for the Ruminant?

During the past 15 years there has been a fascinating array of contributions to understanding the evolution and properties of c-type lysozymes from Wilson, Prager, and their colleagues at the University of California-Berkeley. In the course of their studies, they made a number of interesting observations on stomach lysozymes from ruminants and colobine monkeys (particularly the Hanuman langur, Presbytis entellus).

284

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

Further, they stressed that the convergent evolution of a fermentative foregut in these two groups of mammals has provided them with a unique opportunity to study adaptive evolution at the protein level. In a survey of 23 mammalian species, Dobson et al. (1984) found that c-type lysozyme activity per gram of stomach mucosa is many times higher for ruminants and a leaf-eating (colobinae) monkey than for animals without a foregut. This activity reflects an increased level of lysozyme (molecules) rather than higher specific activity (per molecule). These lysozymes have a narrow pH range of activity, their optimum activities occurring at pH values slightly less than 5.0 for physiological ionic strengths ( I 2 -O.l), in contrast to other c-type lysozymes, whose activity range is broader and optimal at pH -6.5. The ruminant and colobine enzymes are also resistant to pepsin digestion. In light of these and other observations, Dobson et al. (1984) concluded that lysozyme has adapted to function as a digestive enzyme in the true stomach, where it probably degrades the cell walls of bacteria entering from the foregut (for another view see Prieur and Froseth, 1986). Three enzymes (c, , cq, and c Q ) were found to occur in the bovine stomach; the sequence of one of these was determined by Jolles et al. (1984) (see Fig. 10 and Section VII). Subsequently, Stewart et al. (1987) also determined the sequence of the langur stomach lysozyme (see Fig. 10) and further developed the hypothesis of convergent or parallel amino acid replacements (so-called homoplasy). They compared the sequences of the bovine and langur lysozymes with those of the rat, baboon, human, and equine in relation to the biological tree and constructed parsimony trees. If these lysozymes had evolved predominantly by divergence, then the parsimony tree built from them should have matched the branching order of the species. However, the tree placing bovine stomach lysozyme with the langur enzyme was as parsimonious as the biological tree. In the opinion of Stewart et al., homoplasy was the most plausible explanation for this result. This interpretation, involving convergent evolution, and the method of data analysis have been disputed by Cornish-Bowden (1988), and his criticisms were subjected to a spirited reply by Stewart et al. (1988). One feature of the ruminant and Old World monkey stomach lysozymes is their low isoelectric points (pH -7.0-9.0) compared with the high pH value (10.0-12 0’ for many lysozymes. Arg appears to have selected for, during the recent evol.,?‘ :been selected against, ant. ary history of the ruminant lysozymes (Stewart and Wilson, 1987). Lys and Arg are considered generally to be the “epitome of conservative, neutral replacements” (Zuckerkandl and Pauling, 1965; Jukes, 1978).

.

285

LYSOZYME AND ff-LACTALBUMIN

Stewart and Wilson (1987) particularly stressed the lowering of the Argl Lys ratios in these lysozymes. However, the values of Arg/Lys ratios, compared in Table XI, lead us to the conclusion that low ratios are not uniquely associated with the evolution of the ruminant lysozymes. Recently, JolKs et al. (1989) determined amino acid sequences of stomach lysozymes from deer and pig (1,2,3) and compared them with those of stomach lysozymes of bovine ( c 2 ) and langur, and human, baboon, rat, mouse M, chicken, chachalaca, and duck (1) lysozymes. (These sequences are compared in Fig. 10 and Section VII.) They constructed many phylogenetic trees in comparing the sequences of the 13 lysozymes chosen. No tree was (statistically) significantly more parsimonious than the biological tree. They concluded (see their Fig. 4)that the most convergent events took place early in the cow lineage, before it diverged from the deer lineage. The divergence times taken in the comparison were: M (early placental mammals), 60-80 million years ago; A (divergence of pig), 50-60 million years ago; and B (divergence of deer and cow), 20-30 million years ago. The rate of change along AB for lysozyme was considered to be 1.2 changes per million years, or approximately three times the rate for other mammalian lysozymes. In contrast, the average rate from B to present times falls to 0.2 changes per million years, or approximately one-half the “normal” rate for other mammalian lysozymes. Thus, Jolles et al. (1989) confirmed their view that during the period of what they call “recruitment” of lysozyme as a major digestive enzyme in ruminants, the rate of sequence change was accelerated, and later, TABLE XI Comparison of Basic and Acidic Groups and ArglLys Ratios in c-Type Lysozymes No. of residues per molecule

No. of residues per molecule

ArgILys ratio

Species

Arg

Lys

Cow stomach c2 Deer stomach Langur stomach Pig stomach 3 Baboon milk Human milk Horse milk Echidna milk Rat urine Domestic hen egg

3 4 6 7 8 14

11 10 9 13 5

0.29 0.40 0.67 0.54

5

4

15 15 6 6

2.8 0.27 0.2 2.0 1.8

aA

=

3 12 11

1.6

Lys

+ Arg

+

3 4 9 + 6 13 + 7 5 + 8 5 + 14 15+ 4 15+ 3 6 + 12 6 + 11 11

10+

Asp 7 7 9 10 9 8 10 9 9 7

+ Glu + 8 +9 + 3

+3

+ 3 + 3 +6 + 3 + 3 + 2

(Lys + Arg) - (Asp + Glu). Note: Asp and Glu do not include Asn and Gln.

Aa -1

-2 +3 +7 +I +8 +3 +6 +6 +8

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HUGH A. MCKENZIE A N D FREDERICK H. WHITE, JR.

after it was established in its new role, the rate of change was diminished to less than the normal rate. Irwin and Wilson (1989) have shown in a survey of bovine lysozyme clones that lysozyme cp, the most abundant form in bovine stomach mucosa, is encoded by at least two genes, whereas c1 and c3 are possibly encoded by only one gene. They believe that the recruitment involves an early regulatory event followed by a 4- to 7-fold increase in expression allowed by gene amplification. In an interesting paper, Irwin and Wilson (1990) considered the contradictory evolutionary histories of ruminant lysozymes that have been predicted by analysis of genomic blots (Irwin et al., 1989) and sequences of bovine stomach lysozyme cDNAs (Irwin and Wilson, 1989). They characterized stomach lysozyme cDNAs from domestic sheep and axis deer and compared them with one another and with those of the cow. It was found that different parts of the ruminant stomach lysozyme genes have had different evolutionary histories. The 3’ untranslated region has evolved in a divergent fashion since the original duplications 40-50 million years ago, supporting the genomic blot interpretation, whereas the coding region has evolved in a concerted fashion (i.e., the multiple sequences within a species evolved in unison). 4. Divergence of a-Lactalbumin and c-Type Lysozyme

In Section VII,B and Fig. 10 we compared the sequences of 13 alactalbumins (if the bovine A variant, equine B and C variants, and ovine variant are included), 23 mammalian c-type lysozymes (if donkey, mouse M, bovine stomach 1 and 3, caprine 1 and 2, ovine 1-3, camel 1, deer 2, echidna 11, and porcine 1 and 2 are included), and 13 avian c-type lysozymes (if KDIII and PD2 and PD3 are included). Analysis of the sequence differences indicates that, with the recent considerable increase in the number of lysozymes sequenced, there has been an appreciable decrease in the numbers of residues that are invariant in lysozymes as well as for both proteins. Nevertheless, there is still significant overall homology (-35%) between a-lactalbumin and c-type lysozyme. From the similarities in amino acid sequences, three-dimensional structures, intron-exon patterns, etc., there can be little doubt that the concept of divergence is still valid for these proteins. What is controversial are the rate of evolution and the details of the way in which alactalbumin arose, although it is conceded generally that the mechanism involves gene duplication. There are essentially two theories for the divergence, each of which has undergone some modification since it was proposed originally. The first of these, Model I (so called by White et al., 1977, but designated Model I1 in the recent discussion of Prager and Wilson, 1988),

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appears to have originated with Dickerson (see Dickerson, 1971; Dickerson and Geis, 1969). In this model the a-lactalbumin is considered to have arisen about 170-200 million years ago, which Dickerson dated as the start of mammalian development. This duplication is more recent than when those reptiles from which ‘mammallike’ reptiles (and later mammals) diverged from those reptiles whose later divergence produced modern reptiles and birds. With the limited number of a-lactalbumin and mammal and bird lysozyme sequences then available, it appeared that mammalian lysozyme sequences were more similar to those of bird lysozymes than to a-lactalbumins. Hence, Dickerson, in order to make Model I plausible, was led to the proposal that the a-lactalbumin gene underwent accelerated evolution as it acquired its new function and lost the old lysozyme function. As more sequences became available it was evident to Wilson and others that the average rate of sequence change in mammalian lysozymes was virtually the same as that in a-lactalbumin from the time the placental mammals arose. They found that the unit evolutionary period (for a definition see Section X,A,l) for lysozyme was 2.5 X lo6 years and for a-lactalbumin was 2.3 x lo6 years. Thus, Wilson and colleagues (see White et al., 1977; Wilson et al., 1977) were led to propose Model I1 (referred to as Model I by Prager and Wilson, 1988), the essential feature of which is that the a-lactalbumin-lysozyme duplication occurred long before the mammary gland evolved and before the above reptilian split. They believed that this model was in accord with the known sequence resemblances, did not need to invoke rate acceleration, and was, therefore, consistent with the molecular evolutionary clock. In 1984 Brew and colleagues (Shewale et al., 1984) published the sequence of a-lactalbumin from the milk of the red-necked wallaby (Macropus rufogriseus). This paper was of interest in its own right as the first report of the sequence of a marsupial a-lactalbumin, but it was also important in making revisions in the sequences of several other alactalbumins. As a result of the latter, they were able to make proposals for potential binding sites for Ca(I1) to a-lactalbumin (which had recently been shown to be a metalloprotein) and to reassess the evolutionary relationships of a-lactalbumin and lysozyme. Shewale et al. also proposed a modification of Model I, with the gene duplication occurring before the mammary gland arose, but after the reptile split (see Fig. 12). Over the next 2 years (1985- 1986) a number of other developments occurred rapidly. T h e amino acid sequences of echidna lysozymes I and I1 were determined by Teahan et al. (1986, 1991b; see also Griffiths et al., 1985; Teahan, 1986). T h e sequences differed in three residues (Fig. lo),

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but both had several important features: each was (and still is) the only a-lactalbumin or lysozyme that does not have a Cys at position 6, being at 9 instead; and each has all of the residues essential for binding Ca(I1) as in a-lactalbumin (see below), and several other residues normally considered unique to a-lactalbumin. At about the same time Rodriguez et al. (1985) determined and later discussed (Rodriguez et al., 1987) the sequence of pigeon egg-white lysozyme, and, to our surprise, it had residues that indicated potential ability (since confirmed by Nitta et al., 1988) to bind Ca(I1). Further, its sequence terminated at Cys-127, as do echidna lysozyme and wallaby alactalbumin (residue 120, a-lactalbumin numbering). In a sense, a protein such as pigeon lysozyme had been anticipated earlier by White et al. (1977). It is interesting to note that, although lactation is a specific mammalian adaptation, a few birds (e.g., pigeons, emperor penguins, and greater flamingos) secrete a fluid analogous to milk from their gullets. Soon afterward, Stuart et al. (1986; see also Acharya et al., 1989, and Sections V-VII) identified unequivocally the binding sites for Ca(I1) in baboon a-lactalbumin. An examination of Table IX and Fig. 10 indicates that all a-lactalbumins so far sequenced have identical residues in the binding site region, with the exception of rabbit a-lactalbumin, for which residue 79 is Asn instead of Lys and residue 84 is Asn instead of Asp. While it may still be sterically possible for Ca(I1) to be bound in a chelate ring-type arrangement to rabbit a-lactalbumin, it may not be bound as strongly as to other a-lactalbumins. The lack of charge on residues 79 and 84 in the rabbit may be of importance in weakening the binding: Linse et al. (1988) suggested that surface charge may be more important in the binding of metal ions to proteins than is realized generally. As far as we are aware, the extent of binding of Ca(I1) to rabbit a-lactalbumin has not been determined qualitatively nor quantitatively. Of the lysozymes sequenced so far, only echidna has the residues 79, 82, 84, 87, and 88 (a-lactalbumin numbering) in common with a-lactalbumin for Ca(I1) binding, and there is evidence that it does bind Ca(I1) (D. C. Shaw and R. Tellam, personal communication). Equine and pigeon lysozymes have all of the equivalent residues, except for 88, which is Asn instead of Asp. We have already seen in Table XI that both of these lysozymes bind Ca(I1) (Nitta et al., 1987, 1988). Zeng et al. (1990) recently reported the crystallization of equine lysozyme in a form that they consider suitable for X-ray studies. In view of such developments, it is not surprising that there have been several attempts in 1985- 1989 to reconsider the evolutionary relationships. The approach in many studies has been to construct parsimony trees using methods and computer programs based essentially on the maximum parsimony methods of Farris (1970, 1972) or Fitch and Mar-

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goliash (1967). These are distance methods, the starting point being a matrix of pairwise distances, such as the number of amino acid residue differences or minimal mutation distances. In contrast, Prager and Wilson (1988) used character state parsimony analysis. They treated nucleotides as cladistic characters and then looked for shared states at phylogenetically informative sites (for details see page 328 of their paper). An important feature of their work is their attempt to apply tests of statistical significance to the results. Here, we consider briefly three of the recent studies. Prager and Wilson (1988) performed (character state) parsimony analyses of DNA and amino acid sequences for c-type lysozymes of vertebrates and insects and a-lactalbumins. They considered the results presented to provide statistically significant evidence in support of ancient gene duplication, and stated that the period in which this occurred was ‘before the bird-mammal divergence.’ (Presumably, this refers to the type of divergence at A shown in Fig. 12c, one branch of which led to the mammallike reptiles and ultimately the mammals, the other of which later led to divergence of reptiles and birds.) With respect to their model, Prager and Wilson cautiously state, “Although such a demonstration makes this model plausible, we should not consider it established until lactalbumin-like genes (or pseudogenes) or proteins are shown to be present in non mammals.” In the event that the search for lactose and a-lactalbumin in nonmammals is negative, they believe that a long period occurred between gene duplication and the acquisition of specifier activity for lactose synthesis. They included in their analysis only those c-type lysozymes that they call “conventional” and did not include “unconventional” lysozymes (although they presented a justification for this and, at the end of their paper, briefly mentioned the possibility that alactalbumin is related more closely to the unconventional [Ca(II)] binding lysozymes). We believe that preferable terms are “non-calciumbinding” and “calcium-binding” c-type lysozymes, respectively. The Hokkaido group has presented two papers: “The Evolution of Lysozymes and a-Lactalbumin” (Nitta and Sugai, 1989) and “Evolution of Metal Binding Sites in Proteins” (Sugai et nl., 1988). In both studies they included a-lactalbumins and mammalian, avian, and moth (Hyalophora) lysozymes. Nitta and Sugai (1989) examined comprehensively a wide array of evolutionary trees, many of which only prove to be interesting computational exercises. However, some, they concluded, are of evolutionary significance. Their preferred model involves an initial gene duplication before bird-mammal divergence, leading to calcium-binding lysozymes on one lineage and non-calcium-binding lysozymes on another. They then involved a second duplication after the bird-mammal divergence, giving

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rise to mammalian calcium-binding lysozymes in one lineage and, ultimately, a-lactalbumins on the second lineage. They postulated a period of rapid evolution of a-lactalbumin. Three aspects were considered in the development of the model: (1) a-Lactalbumin occurs only in mammals; hence, it is reasonable that it has arisen from a mammalian lysozyme lineage; (2)since Hyulophoru lysozyme does not bind calcium, it is reasonable to assume that the ancestral type did not bind calcium, and if maximum parsimony is observed with respect to acquisition (or loss) of the capacity to bind calcium, the best evolutionary tree groups all of the alactalbumins and calcium-binding lysozymes together; (3) sequence results were analyzed by the Fitch-Margoliash distance method, and also by character state parsimony analysis. Nitta and Sugai (1989) differed from Prager and Wilson (1988) in identifying a period of rapid evolution of a-lactalbumin “not in the process of acquisition of the activity of a-lactalbumin, but after the loss of lysozyme activity.” In both papers the Hokkaido group assumed that echidna lysozyme possesses dual activity and the monotreme lineage is independent of the marsupial and eutherian lineages. These assumptions may not be valid. In the paper by Sugai et al. (1988), it is stated, “The hypothesis of ancient divergence must be abandoned because it is not compatible with a fossil evidence that the lineages of insects and vertebrates diverged long before the radiation of vertebrates.” It is not clear what time scale is involved in this statement. The new studies, especially that by Prager and Wilson (1988), appear to support Model I1 rather than Model I (in the original nomenclature). However, further work is needed to resolve all of the issues involved. C . Are the Functions of Lysozyme and a-Lactalbumin Mutually Exclusive?

The study of the functions of these two proteins is important in gaining an understanding of the evolution of milk proteins and their relationships. Since there seems to have been misunderstanding among some authors about the work on these proteins in the laboratory of one of the authors (H. McK.), we will give a brief history of the findings of these studies to provide clarification. In the early 1960s Mervyn Griffiths, an authority on the biology of monotremes and marsupials, stressed the importance of extending comparative studies to include their milk proteins. Soon after the proposal by Brew and Campbell (1967) of a common ancestor for a-lactalbumin and lysozyme, the studies were extended to include these proteins. It was hoped that the milk of the monotremes, echidna and platypus, might have a primitive a-lactalbumin. Nevertheless, McKenzie was surprised when a graduate student, K. E. Hopper, found a lysozyme in one

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29 1

subspecies of echidna that appeared to have weak specifier activity for the lactose synthase system as well as having lytic activity. By 1970- 1971 preliminary characterization of echidna lysozymes I and 11 had been made (Hopper et al., 1970; Hopper and McKenzie, 1974). No evidence was found for the presence of a “classical”a-lactalbumin. The work was also discussed briefly in a review note, entitled “Milk Proteins in Retrospect and Prospect” (McKenzie, 1971). Hopper and McKenzie (1974) gave details of the lytic activity, weak lactose synthase activity of echidna lysozyme I, the fact that bovine galactosyltransferase could substitute for echidna galactosyltransferase, the isoelectric points of echidna lysozymes I and 11, and the amino acid composition of echidna lysozyme 1. A surprising similarity of echidna lysozyme to equine lysozyme was also noted. Nevertheless, some authors subsequently stated erroneously that alactalbumin, but not lysozyme, was found by Hopper and McKenzie in echidna milk, and it has also been alleged incorrectly that no evidence had ever been presented for the properties of the isolated protein. For logistic reasons, difficulty of locating lactating female monotremes, other urgent priorities, and surprisingly, a lack of financial support and encouragement for this work in Australia (despite its unique fauna), it was not possible to characterize further the echidna lysozymes until the early 1980s. The studies by Teahan et al. (1986, 1990), to which we have already alluded, confirm and extend all but one of the findings of the earlier studies. T h e occurrence of two echidna lysozymes (I and 11) of differing isoelectric points was confirmed. Further, the sequences were found to have unusual features, including certain similarities to equine lysozyme, such as residues for binding Ca(I1). There were weak lactose synthase and lytic activities in the milk samples. However, although the isolated lysozymes possessed lytic activity, they did not act as specifier in the lactose synthase system. The reasons for this are unknown. The obvious possibility is that the original experimental observations of Hopper and McKenzie are in error; however, careful controls were used. A different procedure for isolation was used by Teahan in order to get high yields of lysozyme to enable the determination of the sequence. Commercial galactosyltransferase and a different method were used in the determination of lactose synthase activity of the isolated protein. Nevertheless, it is puzzling, when echidna lysozyme appears to have virtually all of the structural attributes necessary to enable specifier activity, that it was not observed in the recent preparation. This discrepancy and the nature of the specifier protein in the platypus, obviously, must be the subject of further study. It is commonly believed that lactose is the major sugar in milk. However, studies in our laboratory, as well as the more extensive and wideranging studies by Messer and colleagues, indicate that this is not so for

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marsupial and monotreme milk (see, eg., Kamerling et al., 1982; Messer and Green, 1979; Messer and Kerry, 1973; Messer and Mossop, 1977; Messer et al., 1983). For example, echidna milk contains a small amount of free lactose, but much larger amounts of oligosaccharides, such as sialyllactose and fucosyllactose; the major oligosaccharide of platypus milk is 3,2’-difucosyllactose. Thus, either an a-lactalbumin or a lysozyme with specifier activity would be expected in monotreme milk. In addition to the evidence for lactose oligosaccharides in marsupial milk, the occurrence of a-lactalbumin throughout lactation has been demonstrated, for example, in the milk of the red kangaroo [Macropus wfus (Meguleia rufa)] (Bell et al., 1980; McKenzie et al., 1983) and the grey kangaroo (Macropus gzganteus) (McKenzie et al., 1983). An additional a-lactalbumin in the milk of M. gzganteus also appears in late lactation. Lysozyme has also been demonstrated in its milk. These c-type lysozymes and a-lactalbumins have been only partially characterized (see also Brew et al., 1973). T h e only mammal for which the presence of lactose in its milk has not been reported appears to be the California sea lion (Jenness, 1982). No nonmammalian occurrence of a-lactalbumin has been reported. In their theory of the evolution of a-lactalbumin, Hayssen and Blackburn (1985) considered that the duplication of the genetic material for lysozyme occurred as long as 300 million years ago, and that the duplicated material evolved subsequently via an intermediate form with both functions. They also suggested that the protolacteal secretion (from mammary gland precursors) enabled the survival of the eggs or young by virtue of its antimicrobial properties. If the evolution of a-lactalbumin did not occur until after the split of Monotremata and Marsupialia, it is possible that echidna lysozyme could have both functions. [Incidentally, the work by Whittaker et al. (1978) on monotreme hemoglobins and myoglobins does not support a constant evolutionary rate.] It has been conventional wisdom that lysozyme is not active in the lactose synthase system and that a-lactalbumin does not have lytic activity. T h e essential residues for interaction of specifier protein with galactosyltransferase have not yet been unequivocably defined, nor has the role of Ca(I1) in this system. Thus, it is not, at present, possible to rule out weak specifier activity for lysozyme in the lactose synthase system. On the basis of the known structure for c-type lysozyme and the nature of the groups involved in its catalytic activity, and early models of a-lactalbumin structure, there were good structural reasons that militated against a-lactalbumin having lytic activity. This point of view was well developed in the useful reviews by Hill and Brew (1975) and by Brew and Hill (1975). Recent X-ray structural determinations for a-

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lactalbumin by Phillips and co-authors added strength to this point of view (see Acharya et al., 1989), but they did not eliminate the possibility of trace lytic activity for a-lactalbumin and specifier activity for lysozyme. McKenzie and White (1987) studied the question of trace lytic activity in a-lactalbumin. This protein was isolated from a variety of species, and samples were studied for trace lysozyme activity by a sensitive method, previously developed by McKenzie and White ( 1986). All samples exhibof the specific activity of hen eggited trace lytic activity at a level of white lysozyme. Considerable effort was made to avoid contamination by lysozyme in the preparations used. Thus, it was concluded tentatively that the trace lytic activity was due to a-lactalbumin, not to lysozyme contamination. XI. CONCLUSIONS AND

THE

FUTURE

We have tried in this article to integrate the array of investigations on a-lactalbumin and lysozyme. Their range is overwhelming: T h e methods of chemistry, physics, molecular biology, genetics, genetic engineering, comparative biochemistry, enzymology, mathematics, and computer science have been used. Our emphasis has been on the comparative: If we have failed in the integration, at least we hope we have conveyed the spirit of adventure, and that the reader can see that there are further areas to explore. Broadly, there are two aspects in these comparative studies: first, fundamental studies aimed at determining the structure of the proteins, their evolutionary changes, and the structures necessary to maintain their functions and the mechanism of their actions; and second, determination of those biological properties which may lead to their use in pharmacology, medicine, and food science and technology. A tremendous amount has been achieved by the application of X-ray crystallography in determining the three-dimensional structure of alactalbumin and lysozyme, and in the case of lysozyme, the mode of its catalytic action. Nevertheless, despite the tremendous advances, there are still areas of this mechanism that are not fully understood. This is especially true of the comparative mode of action of c-, g-, and phagetype lysozymes. In the case of a-lactalbumin, its mode of interaction with galactosyltransferase and the nature of the critical residues involved have not been clearly defined. The primary and tertiary structures of galactosyltransferase are not known. The mechanism of the catalytic action and the precise role of metal ions are still the subject of disagreement.

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It is evident that there are subtle differences in the structures of c-type lysozymes and a-lactalbumins, beyond what has become evident from Xray crystal structure studies. Scheraga (1988) indicates some of the limitations in current approaches to conformational energy calculations. The procedures used by Scheraga and co-workers for prediction of the protein conformational structure involve a search for the global minimum of the potential energy, allowing additionally for hydration and conformational fluctuational entropy (Gibson and Scheraga, 1987,1988; Kang et al., 1987). The “multiple-minima” problem thus far has hindered computation of the native conformation. However, Scheraga (1988) has made progress in surmounting this problem with peptides u p to 20 residues long. Current efforts are reportedly in progress in studies on peptides of 100-200 residues. Many years ago Linderstr@m-Lang(1952) drew attention to the importance of motility of protein structure. Further work needs to be done on the dynamics of the structures of lysozyme and a-lactalbumin (see also Artymiuk et al., 1979). Kossiakoff (1985) pointed out that the most useful attribute of neutron diffraction studies of proteins (compared with X-ray diffraction) is their ability to locate hydrogen (or deuterium) experimentally. Recent advances in apparatus and data acquisition mean that this method will become increasingly valuable in the study of a-lactalbumin and lysozyme, especially in the location of water molecules and the dynamics of these proteins. An example of a recent application is that by Lehmann et al. (1985). Much of the solution conformational work was done in 1960-1972, when CD, ORD, and Raman spectroscopic apparatus were not very satisfactory. Johnson ( 1988)indicated the scope of secondary structural investigation of proteins with modern CD apparatus. With the increased accessibility of the UV region, such measurements need to be done on a-lactalbumin and lysozyme in a variety of environments over a wide temperature range. Kauzmann (1987) indicated that, in considering the thermodynamics of the unfolding of proteins, we are tending to avoid the hard experiments. Thus, spectroscopic studies of the effects of high pressure on these proteins are sorely needed. Many differences in solution properties between a-lactalbumin and lysozyme are not compatible with prediction and X-ray results, and thus it is surprising that these differences exist. This paradox is consistent with the suggestion by Barman ( 1970), whereby the native tightly folded conformation of a-lactalbumin exists in equilibrium with a “looser” form that is more subject to the various reactions studied, including denaturation. It would also be consistent with the tightly folded but “slowly fluctuating” intermediate of Dolgikh et al. (1981). This equilibrium could

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have contributed to observed difficulties in obtaining crystals suitable for X-ray analysis (until baboon a-lactalbumin was used). There is, moreover, another problem to be considered, assuming the above equilibrium, which is the question of what differences in structural features might permit an equilibrium to exist for a-lactalbumin, while it does not exist for lysozyme. More work is needed to elaborate on this question. For the present it appears that an equilibrium, whereby an alternative and more reactive form of a-lactalbumin is produced, could be a contributing factor in the ability of a-lactalbumin to express traces of cell lytic activity, as well as exhibit the various other unpredictable activities. The use of site-directed mutagenesis in the investigation of a-lactalbumin and lysozyme is in its infancy. Interesting studies are now in progress in the laboratory of A. C. Wilson in collaboration with the laboratory of J. F. Kirsch, and we look forward to future results. Although protein chemists often feel that they now know a good deal about the nature of residues that are critical for proteins to exercise their functions, this may be partially illusory. The study by Luntz et al. (1989) on the structural significance of an internal water molecule studied by sitedirected mutagenesis of Tyr-67 in rat cytochrome c is salutary. While site-directed mutagenesis has a bright future in the study of alactalbumin and lysozyme (see, e.g., Alber et d , 1987; Muraki et d , 1987; Matsumura et al., 1988), this will in no way diminish the need for the study of variants of these proteins from a variety of species. We have stressed in the investigations in our laboratory over the years, and we stress again here without apology, the need to pay special attention to the isolation of these proteins for structural, immunological, and activity studies. There have been unexplained variations in parameters of crystals of a-lactalbumin isolated on separate occasions in some laboratories. Also, the Raman studies by Yu and others [see, e.g., Yu (1977)l indicate appreciable spectral changes on freeze-drying of both a-lactalbumin and lysozyme. We caution against the use of freeze-drying in structural studies. Many of the variants of lysozyme and a-lactalbumin now being studied are available only in small amounts, and there is naturally a temptation to “purify” the protein by HPLC. While this method has considerable advantages, we caution against its use for some purposes. T h e nature of the high pressures involved, and in many cases of the solvents used, can ’bring about irreversible changes. While this may not always be of great importance in amino acid sequence studies, it can be disastrous for conformation and activity studies. Furthermore, many published separations purport to give ‘pure’ peptides and proteins; the nature of the patterns displayed in many publications is testimony to the optimism of the workers involved. Again, we stress that homogeneity cannot be demon-

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strated-only absence of heterogeneity by a particular test. Hence, care should be taken in the choice of such tests, as well as methods of isolation. For both purposes capillary electrophoresis may be the preferred method. It is essential that further work be done on the isolation and characterization of echidna and platypus (monotreme) proteins as well as those of the grey and red kangaroos (marsupials),in order to resolve the issues discussed in Section X. The above remarks on purification are particularly pertinent to such studies. If younger readers are unaware of the traditions of purification of proteins, especially those established in the Carlsberg Laboratory and the former Department of Physical Chemistry at Harvard Medical School, and the rewards for the sheer hard work involved in isolation, we refer them to the recent autobiography of Arthur Kornberg (1989). It has frequently been assumed that lysozymes isolated from different fluids (or organs) from a given species will be identical. Unless the sequences are determined completely, without assumptions from peptide maps, one cannot be certain of this. Again, the elucidation of whether residues are Asp or Asn or Glu or Gln can be dependent on the care taken in treatment of the protein and in sequence determination. The studies by White et al. (1988) on cow milk lysozyme show that the sequence of the milk protein differs from that of bovine stomach lysozyme cp. This indication, that lysozymes from different secretions and tissues of the same species may be different, has been further substantiated by joint studies from two of the most prolific laboratories in lysozyme research: those of Allan Wilson and of Pierre and Jacqueline Jolks. Amino-terminal sequence determinations for caprine stomach lysozymes 1 and 2 and caprine tear lysozymes 1, 2a, and 2b were made by Joll&s et al. (1990). Their results show considerable differences in sequence between the caprine stomach and tear proteins. The 40-residue amino-terminal sequences of the latter bear a striking resemblance to that of cow milk lysozyme. They concluded that their results indicate that the caprine tear family of lysozymes has diverged from the stomach family by an ancient duplication and that later duplications may be responsible for the multiple forms of tear and milk lysozymes in ruminants. Such comparative studies of lysozymes from different secretions and tissues of a given species need to be extended. As well, the genetic association between lysozyme levels in bovine serum and colostrum found by Lie et al. (1986) and the variation in milk found by White et al. (1988) need further study. Great care should be exercised in the isolation of the more labile a-

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lactalbumin if definitive information is to be obtained on the activity of a-lactalbumin in the presence and absence of various metal ions. Above all, the number and nature of metal ion binding sites must be determined and the apparent differences between the results of Phillips’ X-ray group at Oxford University and the views of Kronman (1989) must be resolved. In this connection NMR studies will be very important (see the approach of Adebodun and Jordan, 1989). We have already referred to the considerable discrepancies between values of stability constants from different laboratories. These are summarized in Table XII. The determinations need to be repeated by apTABLE XI1 Association Constantsfor Binding of Ca(II) to a-Lactalbumins and Lysozymes “ ~~

Protein a-Lactalburnin Bovine“

~

Log KsJ 6.4 (log Ks,2 = 4.5 6.4 (pH >6.0)

8.7 9.8 8.0

7.3 9.6

Human Caprine Lysozyme Equine Pigeon

6.9 8.5 8.4 6.3

7.2

Method

Reference

Direct binding, K(I) absent Direct binding, 0.1 M K(I) Fluorometry, chelate presentd Fluorornetry, chelate presentd CD, chelate present: for 25°C by extrapolation Ca electrode, pH 8.0, 37”c CD, chelate and NaCl absent 0.1 M NaCl CD, chelate presentd CD, chelate presentd

1

Dye titration, 0.1 M KC1, 20°C Dye titration, 0.1 M KCI, 20°C

2 3 4 5

6

7 5 5 8 8

“First association constant, Ks,l; second association constant, Kr,2. at 25°C unless indicated. *References: (1) Kronman et al. (1981), ( 2 ) Bratcher and Kronrnan (1984), ( 3 ) Perrnyakov et al. (1981), (4) Murakami et al. (1982),(5) Segawa and Sugai (1983), (6) Hamano et al. (1986), (7) Mitani et al. (1986), ( 8 ) Nitta et al. (1988). “The list for bovine a-lactalbumin is not exhaustive, but is illustrative of the variation in Ks,l values. dChelate is EGTA or EDTA.

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propriate procedures using carefully prepared samples of a-lactalbumin and lysozyme. It is not surprising, because of the antibacterial properties of lysozyme and the variation in levels of occurrence of both a-lactalbumin and lysozyme in various tissues and fluids, that attempts have been made to exploit these properties. An example of this is that lysozyme only occurs at very low levels in cow milk, and hence cow milk-based infant formulas are deficient in this antibacterial agent. Proposals have been made to add domestic hen egg-white lysozyme to boost the level of lysozyme. One of the authors (H. McK.) has advised against this because of its not having identical antigenic properties to human lysozyme and because many commercial samples of hen egg-white lysozyme are contaminated with ovalbumin, a powerful allergen. The revolution in animal breeding in which foreign genes can be substituted for that of lysozyme could be exploited to produce cows that have human lysozyme in their milk (for a general *discussionon the revolution in animal breeding, see Wilmut et al., 1988). Jolles and Jolles (1984) have reviewed the use of lysozyme as a marker in certain diseases. Serum lysozyme levels have been used extensively in the diagnosis of leukemias. Jolles and Jolles discussed some of the reasons for increased and decreased serum levels in various diseases, such as acute or chronic granulocytic leukemia, myeloid metaplasia, and aplastic anemia, and decreased levels in tears in keratoconjunctivitis. They have also considered the interaction of lysozyme with sulfated proteoglycans and its role in the calcification of epiphyseal cartilage. It is to be expected that such studies will yield valuable information, giving rise to further applications in the future (see also Fett et al., 1985). Lysozyme will continue, of course, to serve as a prototype protein for the investigation of the specificity of immune recognition. As Hall and Campbell (1986) have stressed, about one-third of all human metastatic breast carcinomas regress in response to some form of endocrine therapy; yet, despite much research, there is still no reliable way of identifying this group prior to treatment. One approach has been to search for milk proteins, particularly a-lactalbumin, within breast tumors or serum. Despite much effort, Hall and collaborators were unable to find a-lactalbumin being expressed in any of the breast tumors examined (see, e.g., Hall et al., 1981). However, they did find a peptide that was similar, but not identical, to pre-a-lactalbumin. It is to be hoped that the precise nature of the peptide will be determined. Hamilton and collaborators have made extensive studies on the complex processes involved in sperm maturation. We noted in Section VIII that, in the course of this work, they found rat rete testis and epididymal

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fluids to be rich in galactosyltransferase activity. Also, they found an alactalbumin-like protein to be present (see Hamilton, 1981). With a rat mammary gland a-lactalbumin cDNA clone as a hybridization probe, RNA sequences homologous to a-lactalbumin mRNA were detected by Qasba et al. (1983) in the total RNA from rat epididymus. This is taken to mean that a-lactalbumin-like protein is similar in structure to that of a-lactalbumin from the mammary source. In more recent work, De Geyter et al. (1989) suggest that mouse sperm is decapacitated by bovine mammary a-lactalbumin. There is action of a-lactalbumin on the sperm head, inhibiting binding to the zona pellucida. All of this implies a similar function of a-lactalbumin-like protein(s) in the male reproductive tract. It is possible that the a-lactalbumin/lysozyme gene family has a third member. The need for further studies of this member has been stressed in Section VIII. Finally, there are temporal differences in the expression of milk proteins (e.g., casein and a-lactalbumin) among species, exemplified by the rat, guinea pig, and kangaroo (see, e.g., Hall and Campbell, 1986; Burditt et al., 1981). T h e precise causes of these differences remain to be elucidated. ACKNOWLEDGMENTS Work commenced on this article while one of us (H. McK.) was Head of, and the other (F.H.W.) was Visiting Fellow at, the Protein Chemistry Group, John Curtin School of Medical Research, Institute of Advanced Studies, Australian National University, Canberra, ACT 2601, Australia. Warm thanks are expressed to Professors David Phillips and John Edsall for many helpful discussions, and to Dr. Margaret McKenzie for invaluable bibliographical assistance and help. Thanks are due to Drs. Ellen Prager and Allan Wilson for valuable discussions and for generously making available over the years results in advance of publication. H. McK. wishes to thank especially Dr. Mervyn Griffiths for stimulating his interest in the proteins of marsupial and monotreme milk and for invaluable cooperation and help. The skilled help of Panit Thamsongsana in preparing the tables, figures, and manuscript is gratefully acknowledged. One of us (F.H.W.) thanks Robley Light, Chemistry Department, Florida State University, for his generous donation of space and facilities.

REFERENCES Abraham, E. P., and Robinson, R. (1937). Nature (London) 140,24. Acharya, K. R., Stuart, D. I., Walker, N. P. C., Lewis, M., and Phillips, D. C. (1989).J. Mol. Biol. 208,99- 127. Acharya, K. R., Stuart, D. I., Phillips, D. C., McKenzie, H. A., and Teahan, C. G. (1991). Submitted. Achter, E. K., and Swan, I. D. A. (1971). Biochemistry 10,2976-2978. Adebodun, F., and Jordan, F. (1989). Biochemistry 28,7524-7531.

300

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

Alber, T., Dao-pin, S., Wilson, K., Wozniak, J. A., Cook, S. P., and Matthews, B. W. (1987). Nature (London) 330,41-46. Alderton, G., and Fevold, H. L. (1946).J. B i d . C h a . 164, 1-5. Amit, A. G., Mariuzza, R. A., Phillips, S. E. V., and Poljak, R. J. (1986). Sciace 233, 747-753. Andree, P. J., and Berliner, L. J. (1978). Biochim. Biophys. Acta 544,489-495. Andree, P. J., and Berliner, L. J. (1980). Biochemistry 19,929-934. Andrews, A. T. (1983).J. Dairy Res. 50,45-55. Andrews, P. (1969). B i 0 c h . J . 111, 14P-15P. Andrews, P. (1970). FEES Lett. 9,297-300. Anfinsen, C. B. (1959). “The Molecular Basis of Evolution.” Wiley, New York. Anfinsen, C. B., and Scheraga, H. A. (1975). Adv. Protein Chem. 29,205-300. Archer, M., Flannery, T. F., Ritchie, A., and Molnar, R. E. (1985). Nature (London) 318, 363 -366. Armstrong, J. M., McKenzie, H. A., and Sawyer, W. H. (1967). Biochim. Biophys. Acta 147, 60-72. Armstrong, J. M., Hopper, K. E., McKenzie, H. A., and Murphy, W. H. (1970). Biochim. Biophys. Acta 214,419-426. Arnheim, N., Sobel, J., and Canfield, R. (1971).J. Mol. Bzol. 61,237-250. Arnheim, N., Hindenburg, A,, Begg, G. S., and Morgan, F. J. (1973). J. B i d . C h a . 248, 8036-8042. Arnon, R., and Maron, E. (1971).J . Mol. Biol. 61,225-235. Artymiuk, P. J., and Blake, C. C. F. (1981).J. Mol. Biol. 152,737-762. Artymiuk, P. J., Blake, C. C. F., Grace, D. E. P., Oatley, S. J., Phillips, D. C., and Sternberg, M. 1. E. (1979). Nature (London) 280,563-568. Artymiuk, P. J., Blake, C. C. F., Rice, D. W., and Wilson, K. S. (1982). Acta Crystallogr., BS8, 778-783. Aschaffenburg, R., and Drewry, J. (1957). Quoted by Aschaffenburg, R., Occas. Pap., R . Anthropol. Inst. 18,50-54. Aschaffenburg, R., Fenna, R. E., Handford, B. O., and Phillips, D. C. (1972a).J. Mol. B i d . 67,525-528. Aschaffenburg, R., Fenna, R. E., and Phillips, D. C. (1972b).J. Mol. Biol. 67, 529-531. Aschaffenburg, R., Fenna, R. E., Phillips, D. C., Smith, S. G., Buss, D. H., Jenness, R., and Thompson, M. P. (1979).J. Mol. Biol. 127, 135-137. Aschaffenburg, R., Blake, C. C. F., Dickie, H. M., Gayen, S. K., Keegan, R., and Sen, A. (1980). Biochim. Biophys. Actu 625,64-71. Atassi, M. Z.,and Lee, C.-L. (1978). B 2 o c h . J . 171,429-434. Atassi, M. Z.,Habeeb, A. F. S. A., and Rydstedt, L. (1970). Biochim. Biophys. Acta 200, 184- 187. Aune, K. C. (1968). “The Thermodynamics of the Denaturation of Lysozymes,” Ph.D. dissertation. Duke University, Durham, North Carolina. Babad, H., and Hassid, W. Z. (1964).J. B i d . Chem. 239, PC946-PC948. Babad, H., and Hassid, W. 2. (1966).]. B i d . Chem. 241,2672-2678. Bajaj, M., and Blundell, T. (1984). Annu. Rev. Biophys. Bioeng. 13,435-492. Ballardie, F. W., and Capon, B. (1972).J. Chem. SOC.Chem. Commun. pp. 828-829. Banerjee, S. K., Holler, E., Hess, G . P., and Rupley, J. A. (1975). J . B i d . Chem. 250, 4355-4367. Banyard, S. H., Blake, C. C. F., and Swan, I. D. A. (1974). In “Lysozyme” (E. F. Osserman, R. E. Canfield, and S. Beychok, eds.), pp. 71-79. Academic Press, New York. Barel, A. O., Prieels, J.-P., Maes, E., Looze, Y., and Ltonis, J. (1972). Biochim. Biophys. Actu 257,288-296.

LYSOZYME AND CX-LACTALBUMIN

30 1

Barker, R., Olsen, K. W., Shaper, J. H., and Hill, R. L. (1972). J . Biol. Chem. 247, 7135-7147. Barman, T. E. (1970).J . Mol. Biol. 52,391-394. Barman, T. E. (1973). Eur. J . Biochem. 37,86-89. Barman, T. E., and Bagshaw, W. (1972). Biochim. Biophys. Acta 278,491 -500. Baum, J., Dobson, C. M., Evans, P. A,, and Hanley, C. (1989). Biochemistry 28,7-13. Beg, 0.U., von Bahr-Lindstrom, H., Zaidi, Z. H., and Jornvall, H. (1985). Eur. J . Biochem. 147,233-239. Bell, K., and McKenzie, H. A. (1964). Nature (London) 204, 1275-1279. Bell, J. E., Beyer, T. A., and Hill, R. L. (1976).J . Biol. Chem. 251, 3003-3013. Bell, K., Hopper, K. E., McKenzie, H. A., Murphy, W. H., and Shaw, D. C. (1970). Biochim. Biophys. Acta 214,437-444. Bell, K., McKenzie, H. A., Muller, V., and Shaw, D. C. (1980). Mol. Cell. Biochem. 29, 3-8. Bell, K., Hopper, K. E., and McKenzie, H. A. (1981a). A w t . J . Biol. Sci. 34, 149-159. Bell, K., McKenzie, H. A., Muller, V., Rogers, C., and Shaw, D. C. (1981b). Comp. Biochem. Physiol. 68B, 225-236. Bell, K., McKenzie, H. A., and Shaw, D. C. (1981~). Mol. Cell. Biochem. 35, 113-119. Benjamin, D. C., Berzofsky, J. A., East, I. J., Curd, F. R. N., Hannum, C., Leach, S. J., Margoliash, E., Michael, J. G., Miller, A., Prager, E. M., Reichlin, M., Sercarz, E. E., Smith-Gill, S. J., Todd, P. E., and Wilson, A. C. (1984). Annu. Rev. Immunol. 2, 67-101. Benton, M. (1984). New Sci. 103, 18-19. Benton, M. J. (1990).J . Mol. Evol. 30, 409-424. Berger, L. R., and Weiser, R. S. (1957). Biochim. Biophys. Acta 26,517-521. Berliner, L. J., and Kaptein, R. (1981). Biochemistry 20,799-807. Berliner, L. J., and Johnson, J. D. (1988). In “Calcium Binding Proteins” (M. P. Thompson, ed.), Vol. 11, pp. 79- 116. CRC Press, Boca Raton, Florida. Berliner, L. J., and Robinson, R. D. (1982). Bzochemist7y 21,6340-6343. Berliner, L. J., Ellis, P. D., and Murakami, K. (1983). Biochemistry 22,5061-5063. Berliner, L. J., Davis, M. E., Ebner, K. E., Beyer, T. A., and Bell, J. E. (1984). Mol. Cell. Biochem. 62,37-42. Berthou, J., and Jolles, P. (1974). Biochim. Biophys. Acta 336,222-227. Berthou, J., Lifchitz, P., Artymiuk, P., and Jolles, P. (1983). Proc. R. SOC.London, Ser. B 217, 47 1-489. Bhattacharjee, N., and Vonderhaar, B. K. (1983). Biochim. Biophys. Acta 755,279-286. Blake, C. C. F. (1983). TrendcBzochem. Sci. (Pers. Ed.) 8, 11-13. Blake, C. C. F., Fenna, R. H., North, A. C. T., Phillips, D. C., and Poljak, R. J. (1962). Nature (London) 196, 1173- 11’76. Blake, C. C. F., Koenig, D. F., Mair, G. A., North, A. C. T., Phillips, D. C., and Sarma, V. R. (1965). Nature (London) 206,757-761. Blake, C. C. F., Mair, G. A., North, A. C. T., Phillips, D. C., and Sarma, V. R. (1967a). Proc. R. SOC.London, Ser. B 167,365-377. Blake, C. C. F., Johnson, L. N., Mair, G. A., North, A. C. T., Phillips, D. C., and Sarma, V. R. (1967b). Proc. R . SOC.London, Ser. B 167,378-388. Blake, C. C. F., Pulford, W. C. A., and Artymiuk, P. J. (1983).J . Mol. Biol. 167,693-723. Bloomfield, A. (1919). Bull. Johns Hopkins Hosp. 30,317-322. Bloomfield, A. (1920). Bull, Johns Hopkim Hosp. 31, 14-19, 85-89, and 203-206. Blumberg, B. S., and Tombs, M. P. (1958). Nature (London) 181,683. Blundell, T. L., and Johnson, L. N. (1976). “Protein Crystallography.” Academic Press, New York. Boesman-Finkelstein, M., and Finkelstein, R. A. (1982). FEBS Lett. 144, 1-5.

302

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

Bordet, M. J., and Bordet, M. (1924). C . R. Hebd. Skances Acad. Sci. 179, 1109- 1 1 13. Bradbury, J. H., and King, N. L. R. (1971). Azcst.J. Chem. 24, 1703-1714. Bradbury, J. H., and Norton, R. S. (1975). Eur.J. Biochem. 53,387-396. Bragg, L. (1967). Proc. R. SOC.London, Ser. B 167,349. Bratcher, S. C., and Kronman, M. J. (1984).J. Biol. Chem. 259, 10875-10886. Brew, K. (1969). Nature (London) 222,671-672. Brew, K. (1970). Essays B i o c h . 6,93- 118. Brew, K. (1972). Eur.J. Biochem. 27,341-353. Brew, K., and Campbell, P. N. (1967). Bzochem.J. 102,258-264. Brew, K., and Hill, R. L. (1975). Rev. Physiol. B z o c h . P h m o l . 72, 105-158. Brew, K., Vanaman, T. C., and Hill, R. L. (1967).J. Biol. C h . 242,3747-3749. Brew, K., Vanaman, T. C., and Hill, R. L. (1968). Proc. Natl. Acad. Sci. U.S.A. 59,491-497. Brew, K., Castellino, F. J., Vanaman, T. C., and Hill, R. L. (1970). J . Biol. Chem. 245, 4570-4582. Brew, K., Steinman, H. M., and Hill, R. L. (1973).J.Biol. Chem. 248,4739-4742. Brodbeck, U., and Ebner, K. E. (1966).J.Bzol. Chem. 241,762-764. Brodbeck, U., Denton, W. L., Tanahashi, N., and Ebner, K. E. (1967).J . Biol. Chem. 242, 1391- 1397. Brooks, C. L., 111, and Karplus, M. (1989).J. Mol. Biol. 208, 159-181. Brown, J. R. (1964). Biochem.J. 92, 13 p. Brown, R. C., Fish, W. W., Hudson, B. G., and Ebner, K. E. (1977). Bzochim. Bzophys. Acta 441,82-92. Browne, W. J., North, A. C. T., Phillips, D. C., Brew, K., Vanaman, T. C., and Hill, R. L. (1969).J . Mol. BWl. 42,65-86. Brusca, A. E., and Patrono, D. (1960). G. Batteriol., Vzrol. Zmmunol. 53, 211-214. Burditt, L. J., Parker, D., Craig, R. K., Getova, T., and Campbell, P. N. (1981). Bi0chem.J. 194,999- 1006. Campbell, K. S. W., and Day, M. F. (1987). In “Rates of Evolution” (K. S. W. Campbell and M. F. Day, eds.), p. vii. Allen & Unwin, London. Canfield, R. E. (1963).J . Biol. Chem. 238,2698-2707. Canfield, R. E., and Liu, A. K. (1965).J. Biol. Chem. 240, 1997-2002. Canfield, R. E., and McMurry, S. (1967). Bzochem. Biophys. Res. Commun. 26, 38-42. Canfield, R. E., Kammerman, S., Sobel, J. H., and Morgan, F. J. (1971). Nature (London), New Biol. 232, 16- 17. Canfield, R. E., Collins, J. C., and Sobel, J. H. (1974). In “Lysozyme” (E. F. Osserman, R. E. Canfield, and S. Beychok, eds.), pp. 63-70. Academic Press, New York. Careri, G., Gratton, E., Yang, P.-H., and Rupley, J. A. (1980). Nature (London) 284, 572-573. Carr, C. W. (1953). Arch. Bzochem. Biophys. 46,424-431. Carroll, R. L. (1988). “Vertebrate Paleontology and Evolution.” Freeman, New York. Castafion, M. J., Spevak, W., Adolph, G. R.,Chlebowicz-Sledziewska,E., and Sledziewski, A. (1988). Gene 66,223-234. Castellino, F. J., and Hill, R. L. (1970).J.Biol. Chem. 245,417-424. Castellino, F. J., Fish, W. W., and Mann, K. G. (1970).J.Biol. Chem. 245,4269-4275. Chandan, R. C., Parry, R. M., and Shahani, K. M. (1965). Bzochzm. Biophys. A c h 110, 389-398. Chandler, D. K., Silvia,J. C., and Ebner, K. E. (1980). Bzochim. Bzophys. Acta 616, 179-187. Chen, M. C., Lord, R. C., and Mendelsohn, R. (1974).J. Am. Chem. SOC.96, 3038-3042. Chen, Y.-H., Yang, J. T., and Chau, K. H. (1974). Biochemistry 13,3350-3359. Chiarandra, G., Saccomani, G., and Tamburro, A. M. (1972). Experientia 28, 1405-1406.

LYSOZYME AND (Y-LACTALBUMIN

303

Chou, P. Y., and Fasman, G. D. (1974). Biochemistry 13,222-245. Chung, L. P., Keshav, S., and Gordon, S. (1988). Proc. Natl. Acud. Sci. U.S.A. 85, 6227-623 1. Clemens, W. A. (1989). I n “Fauna of Australia” (D. W. Walton and B. J. Richardson, eds.), Vol. IB, pp. 401-406. Aust. Govt. Publ. Serv., Canberra, Australia. Clymer, D. C., Geren, C. R., and Ebner, K. E. (1976). Biochemistry 15, 1093-1097. Conti, A., Godovac-Zimmermann, J., Napolitano, L., and Liberatori, J. (1985). MiZchwGsenschaft 40,673-675. Cornish-Bowden, A. (1979). J. Theor. Biol. 76,369-386. Cornish-Bowden, A. (1988). Nature (London) 332,787-788. Cortopassi, G. A., and Wilson, A. C. (1989). I n “The Immune Response to Structurally Defined Proteins: T h e Lysozyme Model” (S. Smith-Gill and E. Sercarz, eds.), pp. 87-96. Adenine Press, Schenectady, New York. Cowburn, D. A., Bradbury, E. M., Crane-Robinson, C., and Gratner, W. B. (1970). Eur. J. Biochem. 14, 83-93. Cowburn, D. A,, Brew, K., and Gratzer, W. B. (1972). Biochemistry 11, 1228-1234. Craig, R. K., Hall, L., Parker, D., and Campbell, P. N. (1981). E i o c h a . J. 194, 989-998. Creighton, T. E. (1984). “Proteins: Structures and Molecular Principles.” Freeman, New York. Crompton, A. W. (1968). Optima 18, 137-151. Cross, M., Mangelsdorf, I., Wedel, A., and Renkawitz, R. (1988). PrQc. Natl. Acad. Scz. U.S.A. 85, 6232-6236. Cunningham, W. P., Mollenhauer, H. H., and Nyquist, S. E. (1971). J . Cell Biol. 51, 273-285. Dandekar, A. M., and Qasba, P. K. (1981). Proc. Natl. Acad. Sci. U.S.A. 78,4853-4857. Dao-pin, S., Liao, D.-I., and Remington, S. J. (1989). Proc. Natl. Acud. Sci. U.S.A. 86, 5361 -5365. Dawson, T. J. (1977). Sci. Am. 237,78-89. De Geyter, C., Cooper, T. G., De Geyter, M., and Nieschlag, E. (1989). Gamete Res. 24, 415-426. Deonier, R. C., and Williams, J. W. (1970). Biochemistry 9,4260-4267. Desmet, J., Hanssens, I., and Van Cauwelaert, F. (1987). Biochim. Biophys. Acta 912, 211-219. Desmet, J., Van Dael, H., Van Cauewelaert, F., Nitta, K., and Sugai, S. (1989). J . Znorg. Biochem. 37, 185-191. Dianoux, A. C., and Joll& P. (1967). Biochim. Biophys. Actu 133,472-479. Dickerson, R. E. (1971).J. Mol. Evol. 1, 26-45. Dickerson, R. E., and Geis, I. (1969). “The Structure and Action of Proteins,” pp. 69-78. Harper & Row, New York. Dickman, S. R., and Proctor, C. M. (1952). Anal. Biochem. Biophys. 40, 364-372. Dobson, C. M., and Williams, R. J. P. (1977). In “Metal-Ligand Interactions in Organic Chemistry and Biochemistry” (B. Pullman and N. Goldblum, eds.), Part 1, pp. 255282. Reidel, Dordrecht, T h e Netherlands. Dobson, D. E., Prager, E. M., and Wilson, A. C. (1984).J.Biol. Chem. 259, 11607-11616. Dolgikh, D. A,, Gil’manshin, R. J., Brazhnikov, E. V., Bychkova, V. E., Semisotnov, G. V., Venyaminov, S. Y., and Ptitsyn, 0. B. (1981). FEBS Lett. 136,311-315. Dolgikh, D. A., Abaturov, L. U., Bolotina, I. A., Brazhnikov, E. V., Bychkova, V. E., Gil’manshin, R. I., Lebedev, Y. O., Semisotnov, G. V., Tiktopulo, E. I., and Ptitsyn, 0. B. (1985). Eur.Eiophys.J . 13, 109-121. Douzou, P. (1977). “Cryobiochemistry,” pp. 236-267. Academic Press, New York.

304

H U G H A. MCKENZIE AND FREDERICK H. WHITE, JR.

Douzou, P., Hoa, G. H. B., and Petsko, G. A. (1974).J.Mol. B i d . 96,367-380. Ebner, K. E. (1970). Acc. Chem. Res. 3,41-47. Ebner, K. E., Denton, W. L., and Brodbeck, U. (1966). Bzocha. Biophys. Res. Commun. 24, 232-236. Edsall, J. T. (1938). Cold Spring Harbor Symp. Quant. Biol. 6,40-49. Edsall, J. T., and McKenzie, H. A. (1983). Adv. Biophys. 16, 53-183. Ehrenpreis, S., and Warner, R. C. (1956). Arch. Biochem. Biophys. 61,38-50. Ekstrand, B., and Bjorck, L. (1986).J.Chromatogr. 358,429-433. Engstrom, Xanthopoulos, K. G., Boman, H. G., and Bennich, H. (1985). EMBOJ. 4, 2119-2122. Epstein, C. A., and Chain, E. (1940). Br. J . Exp. Pathol. 2,l, 339-355. Fainanu, M., Wilson, A. C., and Arnon, R. (1974).J. MoE. B i d . 84,635-642. Farris, J. S. (1970). Syst. Zool. 19, 83-92. Farris, J. S. (1972). Am. Nat. 106,645-668. Faure, A., and Jolles, P. (1970). FEBS Lett. 10,237-240. Fenna, R. E. (1982a).J. Mol. B i d . 161,203-210. Fenna, R. E. (1982b).J. MoLBiol. 161,211-215. Fernandez-Sousa, J. M., Gavilanes,J. G., Municio, A. M., and Rodriguez, R. (1979). Comp. Biochem. Physiol. 62B, 389-392. Fett, J. W., Strydom, D. J., Lobb, R. R., Alderman, E. M., and Vallee, B. L. (1985).Biochemistry 24,965-975. Fiess, H. A., and Klotz, I. M. (1952).J.Am. Chem. SOC.74,887-891. Findlay, J. B. C., and Brew, K. (1972). Eur. J. Biochem. 27,65-86. Fitch, W. M., and Margoliash, E. (1967). Science 155,279-284. Fitzgerald, D. K., Brodbeck, U., Kiyosawa, I., Mawal, R., Colvin, B., and Ebner, K. E. (1970a).J. B i d . Chem. 245,2103-2108. Fitzgerald, D. K., Colvin, B., Mawal, R., and Ebner, K. E. (1970b). Anal. B i o c h a . 36, 43-61. Fleming, A. (1922). Proc. R. Sac. London, Ser. B 93, 306-317. Fleming, A. (1932). Proc. R . SOC.Med. 26,71-84. Fleming, A., and Allison, V. D. (1922). Br. J. Exp. Pathol. 3, 252-260. Fokker, A. P. (1890). Fwtschr. Med. 8,7-8. Ford, L. O . ,Johnson, L. N., Machin, P. A., Phillips, D. C., and Tjian, R. (1974).J.Mol. Biol. 88,349-371. Fouche, P. B., and Hash, J. H. (1978).J. B i d . Chem. 253,6787-6793. Fraser, I. H., and Mookerjea, S. (1976). Biochem.J. 156,347-355. Fraser, I. H., Wadden, P., and Mookerjea, S. (1980). Can.J. Biochem. 58,878-884. Friedberg, F. (1988). B i o c k . E d w . 16,35-36. Galat, A., Creighton, T. E., Lord, R. C., and Blout, E. R. (1981). Biochemistry 20,594-601. Gallo, A. A,, Swift, T. J., and Sable, H. Z. (1971). Biochem. Biophys. Res. Commun. 43, 1232- 1238. Gardiner, B. G. (1982). Zool. J. Linn. SOC.74,207-232. Garfinkel, D., and Edsall, J. T. (1958).J.Am. C h a . SOC.80,3818-3823. Gauthier, J., Kluge, A. G., and Rowe, T. (1988). Cladistics 4, 105-209. Gavilanes,J. G., MenCndez-Arias, L., and Rodriguez, R. (1984). Comp. Biochem. Physiol. 77B, 83-88. Gaye, P., Hue-Delahaie, D., Mercier, J.-C., Soulier, S., Vilotte,J.-L., and Furet, J.-P. (1987). Biochimie 69,60 1-608. Giblett, E. R. (1969). “Genetic Markers in Human Blood.” Blackwell, Oxford, England.

w.,

Gibson, K. D., and Scheraga, H. A. (1987).J . Cornput. Chon. 8,826-834.

LYSOZYME A N D ff-LACTALBUMIN

305

Gibson, K. D., and Scheraga, H. A. (1988). In “Structure and Expression” (M. H. Sarma and R. H. Sarma, eds.), Vol. 1 , pp. 67-94. Adenine Press, Schenectady, New York. Gilbert, W. (1978). Nature (London), 271,501. Gil’manshin, R. I., Ptitsyn, 0. B., and Semisotnov, G. V. (1988). Biofiziku 33, 217-221. Glazer, A. N., and Simmons, N. S. (1965).J. Am. Chem. Soc. 87,2287-2288. Godovac-Zimmermann, J., Shaw, D. C., Conti, A., and McKenzie, H. A. (1987). B i d . Chem. Hoppe-Seyler 368,427-433. Godovac-Zimmermann, J., Conti, A., and Napolitano, L. (1988). Biol. Chem. Hoppe-Seyler 369,1109-1115. Gordon, W. G. (1971). In “Milk Proteins” (H. A. McKenzie, ed.), Vol. 11, pp. 331-365. Academic Press, New York. Greenfield, N., and Fasman, G. C. (1969). Biochemistry 8,4108-41 16. Griffiths, M. (1978). “The Biology of the Monotremes.” Academic Press, New York. Griffiths, M. (1988). Sci. Am. 258,84-91. Griffiths, M. (1989). In “Fauna of Australia” (D. W. Walton and B. J. Richardson, eds.), Vol. IB, pp. 407-435. Aust. Govt. Publ. Serv., Canberra, Australia. Griffiths, M. E., McKenzie, H. A., Shaw, D. C., and Teahan, C. G. (1985). Proc. A w t . Biochem. Soc. 17, 25. Grinde, B., Jollhs, J., and Joll6s, P. (1988). Eur.J. Biochem. 173,269-273. Gronenborn, A. M., and Clore, G. M. (1990). Anal. Chem. 62,2-15. Grosclaude, F., Mahk, M.-F., Mercier, J. C., Bonnemaire, J., and Tessier, J. H. (1976). Anim. Genet. Sel. Anim. 8,461-479. Grunwald, J., Dhawale, S. W., and Berliner, L. J. (1982). Int. J. B i d . Macromol. 4 , 3 7 1-374. Griitter, M. G., Weaver, L. H., and Matthews, B. W. (1983). Nature (London) 303,828-831. Hall, L., and Campbell, P. N. (1986). EssuysBiochem. 22, 1-26. Hall, L., Davies, M. S., and Craig, R. K. (1981). Nucleic Acids Res. 9,65-84. Hall, L., Craig, R. K., Edbrooke, M. R., and Campbell, P. N. (1982). Nucleic Acids Res. 10, 3503-3515. Hall, L., Emery, D. C., Davies, M. S., Parker, D., and Craig, R. K. (1987). Bzochem.]. 242, 735-742. Halliday, J. A,, Bell, K., McKenzie, H. A., and Shaw, D. C. (1990). Comp. Biochem. Physiol. 95B, 773-779. Halper, J. P., Latovitzki, N., Bernstein, H., and Beychok, S. (1971). Proc. Natl. Acad. Sci. U.S.A. 68,517-522. Hamano, M., Nitta, K., Kuwajima, K., and Sugai, S. (1986). /. Biochem. (Tokyo) 100, 1617-1622. Hamilton, D. W. (1980). Biol. Reprod. 23, 377-385. Hamilton, D. W. (1981). B i d . Reprod. 25, 385-392. Hammer, M. F., and Wilson, A. C. (1990). Manuscript in preparation. Hanson, L. A. (1960). Int. Arch. Allergy Appl. Immunol. 17,45-69. Hanssens, I., Pottel, H., Herreman, W., Van Cauwelaert, F. (1984). Biochem. Biophys. Res. Commun. 119,509-515. Harada, S., Sarma, R., Kakudo, M., Hara, S., and Ikenaka, T. (1981).J. Biol. Chem. 256, 11600- 11602. Hartley, B. S., Brown, J. R., Kauffman, D. L., and Smillie, L. B. (1965). Nature (London) 207,1157- 1159. Hayashi, K., Imoto, T., and Funatsu, M. (1964).J. Biochem. (Tokyo) 55,516-521. Hayssen, V., and Blackburn, D. G. (1985). Evolution 39, 1147- 1149. Hermann, J., and Jolles, J. (1970). Biochim. Biqhys. Acta 200, 178-179. Hermann, J., Jollhs, J., and Joll*s, P. (1971). Eur.J. Biochem. 24, 12-17.

306

HUGH A. MCKENZIE AND FREDERICK H . WHITE, JR.

Hermann, J., Joll&s,J., Buss, D. H., and Jollks, P. (1973). J . Mol. B i d . 79, 587-595. Hill, R. L., and Brew, K. (1975). Adv. Enzyme Relat. Areas Mol. B i d . 43,411-490. Hill, R. L., Delaney, R., Fellows, R. E., Jr., and Lebovitz, H. E. (1966). Proc. Natl. Acad. Sci. U.S.A. 56, 1762-1769. Hill, R. L., Brew, K., Vanaman, T. C., Trayer, I. P., and Mattock, P. (1969). Brookhaven SymP. B i d . 21, 139-154. Hill, R. L., Steinman, H. M., and Brew, K. (1974). I n “Lysozyme” (E. F. Osserman, R. E. Canfield, and S. Beychok, eds.), pp. 55-62. Academic Press, New York. Hiraoka, Y., and Sugai, S. (1984). Int. J . Pept. Protein Res. 23,535-542. Hiraoka, Y., and Sugai, S. (1985). Int. J. Pept. Protein Res. 26,252-261. Hiraoka, Y., Segawa, T., Kuwajima, K., Sugai, S., and Murai, N. (1980). Biochem. Biophys. Res. Commun. 95, 1098- 1194. Hogle, J., Rao, S. T., Mallikarjunan, M., Beddell, C., McMullan, R. K., and Sundaralingam, M. (1981). Acta Crystallogr. B37,591-597. Holladay, L. A,, and Sophianopoulos, A. J. (1972).J. Biol. Chem. 247, 1976-1979. Hopp, T. P., and Woods, K. R. (1979). Biochemistry 18,5182-5191. Hopp, T. P., and Woods, K. R. (1982). Mol. Immunol. 19, 1453-1463. Hopper, K. E. (1973). Biochim. Biophys. Acta 295,364-370. Hopper, K. E., and McKenzie, H. A. (1973). Biochim. Biophys. Acta 295,352-363. Hopper, K. E., and McKenzie, H. A. (1974). Mol. Cell. Biochem. 3,93-108. Hopper, K. E., McKenzie, H. A., and Treacy, G. B. (1970). Proc. A w t . Biochem. SOC.3, 86. Howard, J. B., and Glazer, A. N. (1969)./. Biol. Chem. 244,1399- 1409. Hurley, W. L., and Schuler, L. A. (1987). Gene 61, 119-122. Hymes, A. J., and Mullinax, F. (1984). Anal. Biochem. 139,68-72. Ibrahimi, I. M., Prager, E. M., White, T. J., and Wilson, A. C. (1979). Biochemistry 18, 2736-2744. Ikeda, K., and Hamaguchi, K. (1973).J.Biochem. (Tokyo)73,307-322. Ikeguchi, M., Kuwajima, K., and Sugai, S. (1986).J. Biochem. (Tokyo)99, 1191-1201. Imoto, T. Johnson, L. N., North, A. C. T., Phillips, D. C., and Rupley, J. A. (1972). I n “The Enzymes” (P. D. Boyer, ed.), 3rd Ed., Vol. VII, pp. 665-868. Academic Press, New York. lmoto, T., Andrews, L. J., Banerjee, S. K., Shrake, A., Forster, L. S., and Rupley, J. A. (1975).J . B i d . Chem. 250,8275-8282. Imoto, T., Ono, T., and Yamada, H. (1981).J. Biochem. (Tokyo)90, 335-340. Ingram, V. M. (1963). “The Hemoglobins in Genetics and Evolution.” Columbia Univ. Press, New York. Irwin, D. M., and Wilson, A. C . (1989).J. Biol. Chem. 264, 11387- 11393. Irwin, D. M., and Wilson, A. C. (1990).J . B i d . Chem. 265,4944-4952. Irwin, D. M., Sidow, A., White, R. T., and Wilson, A. C. (1989). In “The Immune Response to Structurally Defined Proteins: The Lysozyme Model” (S. Smith-Gill and E. Sercarz, eds.), pp. 73-85. Adenine Press, Schenectady, New York. Isaacs, N. W., Machin, K. J., and Masakuni, M. (1985). Azcst.J. B i d . Sci. 38, 13-32. Iyer, K. S., and Klee, W. A. (1973).J. Biol. Chem. 248,707-710. Jenness, R. (1982). In “Developments in Dairy Chemistry” (P. F. Fox, ed.), Vol. 1, pp. 87114. Elsevier, London. Johnson, L. N., and Phillips, D. C . (1965). Nature (London) 206,761-763. Johnson, L. N., Cheetham, J., McLaughlin, P. J., Acharya, K. R., Barford, D., and Phillips, D. C. (1988). CUT. Top.Microbiol. Immunol. 139,s 1 - 134. Johnson, W. C., Jr. (1988). Annu. Rev. Biophys. Biophys. Chem. 17, 145-166.

LYSOZYME AND (Y-LACTALBUMIN

307

Jolles, J., and Jollts, P. (1971). Helv. Chirn. Actu 54, 2668-2675. Jolles, J., and Jolles, P. (1972). FEBS Lett. 22, 31-33. Jollks, J., Jhuregui-Adell, J., Bernier, I., and Jollks, P. (1963). Biochim. Biophys. Acta 78, 668-689. Jollks, J., Jauregui-Adell, J., and Jollts, P. (1964). C . R. Hebd. Siunces Acad. Sci. 258, 3926-3928. Jollts, J., Van Leemputten, E., Mouton, A., and Jolles, P. (1972). Biochim. Biophys. Actu 257, 497-5 10. Jolles, J., Schoentgen, F., Jolles, P., Prager, E. M., and Wilson, A. C. (1976). J . Mol. Evol. 8, 59-78. Jolles, J., Ibrahimi, I. M., Prager, E. M., Schoentgen, F., Jollks, P., and Wilson, A. C. (1979a). Biochemistry 18,2744-2752. Jolles, J., Schoentgen, F., Croizier, G., Croizier, L., and Jolles, P. (1979b). J , Mol. Evol. 14, 267-27 1. Jollks, J., Jolles, P., Bowman, B. H., Prager, E. M.,Stewart, C.-B., and Wilson, A. C. (1989). J . Mol. Evol. 28,528-535. Jolles, J., Prager, E. M., Alnemri, E. S., Jolles, P., Ibrahimi, I. M., and Wilson, A. C. (1990). J . Mol. E d . 30, 370-382. Jolles, P., and Berthou, J. (1972). FEBS Lett. 23, 21-23. Jollts, P., and Fromageot, C. (1954). Biochim. Biophys. Acta 14, 219-227. Jolles, P., and Jolles, J. (1984). Mol. Cell. Biochem. 63, 165-189. Jolles, P., and Ledieu, M. (1959). Biochim. Biophys. Acta 31, 100-103. Jollts, P., Schoentgen, F., Jolles, J., Dobson, D. E., Prager, E. M., and Wilson, A. C. (1984). J . Biol. Chem. 259, 11617- 11625. Jones, R., Dwek, R. A., and Forsen, S. (1974). Eur. J . Biochem. 47,271-283. Jori, G., Gennari, G., Galiazzo, G., and Scoffone, E. (1971). Biochirn. Biophys. Acta 236, 749-766. Joynson, M. A,, North, A. C. T., Sarma, V., Dickerson, R. E., and Steinrauf, L. K. (1970). J. Mol. Biol. 50, 137-142. Jukes, T. H. (1978). Adv. Enzymol. 47,375-432. Jung, A,, Sippel, A. E., Grez, M., and Schutz, G. (1980). Proc. Natl. Acud. Sci. U.S.A. 77, 5759-5763. Kamerling, J. P., Dorland, L., van Halbeek, H., Vliegenthart, J. F. G., Messer, M., and Schauer, R. (1982). Curbohydr. Res. 100,331 -340. Kaminogawa, S., Mizobuchi, H., and Yamaguchi, K. (1972). Agrzc. Biol. Chem. 12, 2 163-2 167. Kaminogawa, S . , McKenzie, H. A., and Shaw, D. C. (1982). Proc. 1nt. Congr. Biochem., 12th p. 325. Kaminogawa, S., McKenzie, H. A., and Shaw, D. C. (1984). Biochem. Int. 9,539-546. Kaneda, M., Kato, I., Tominaga, N., Titani, K., and Narita, K. (1969). J . Biochern. (Tokyo) 66,747-749. Kang, Y. K., NCmethy, G., and Scheraga, H. A. (1987). J. Phys. Chem. 91, 4105-4109, 4109-4118, and 4118-4120. Kato, S., Okamura, M., Shimamoto, N., and Utiyama, H. (1981). Biochemistry 20, 1080-1085. Kato, S . , Shimamoto, N., and Utiyama, H. (1984). Biochim. Biophys. Actu 784, 124-132. Kauzmann, W. (1987). Nature (London) 325,763-764. Kelly, J. A., Sielecki, A. R., Sykes, B. D., James, M. N. G , and Phillips, D. C. (1979). Nature (London) 282,875-878.

308

HUGH A. MCKENZIE A N D FREDERICK H. WHITE, JR.

Kemp, T. S. (1983).J . Linn. SOC.77,353-384. Kerby, G. P., and Eadie, G. S. (1953).Proc. SOC.Exfi. Biol. Med. 83, 1 1 1 - 113. Khatra, B. S.,Herries, D. G., and Brew, K. (1974).Eur. J. Biochem. 44,537-560. Kimura, M. (1983).“The Neutral Theory of Molecular Evolution.” Cambridge Univ. Press, Cambridge, England. Kimura, M. (1987).J.Mol. Evol. 26,24-33. Kirsch, J. F., Malcolm, B. A., Corey, M. J., Zhang, L., and Wilson, A. C. (1989).I n “The Immune Response to Structurally Defined Proteins: The Lysozyme Model” (S. SmithGill and E. Sercarz, eds.), pp. 59-63. Adenine Press, Schenectady, New York. Kim, N., Kuwajima, K., Nitta, K., and Sugai, S. (1976).Biochim. Biophys. Actu 427,350-358. Kitchen, B.J., and Andrews, P. (1974).Bi0chem.J. 141, 173-178. Klee, W. A., and Klee, C. B. (1970).Biochem. Biophys. Res. Commun. 39, 833-841. Klee, W. A., and Klee, C. B. (1972).J. Bzol. Chem.247,2336-2344. Klotz, I. M. (1970).Arch. Biochem. Biophys. 138, 704-706. Koga, K., and Berliner, L. J. (1985).Biochemistry 24,7257-7262. Kondo, K., Fujio, H., and Amano, T. (1982).J. Biochem. (Tokyo) 91, 571-587. Kornberg, A. (1989).“For the Love of Enzymes: The Odyssey of a Biochemist,” Harvard Univ. Press, Cambridge, Massachusetts. Kossiakoff, A. A. (1985).Annu. Rev. Biochem. 54, 1195-1227. Kretsinger, R. H. (1976).Int. Rev. Cytol. 46,323-393. Krigbaum, W. R.,and Kiigler, F. R. (1970).Biochemistry 9, 1216-1223. Kronman, M. J. (1968).Biochem. BZSphys. Res. Commun. 33,535-541. Kronman, M. J. (1989).CRC Crit. Rev. Biochem. 24,565-667. Kronman, M. J., and Andreotti, R. (1964).Biochemistry 3, 1145-1 151. Kronman, M. J., and Bratcher, S. C. (1983).J.Biol. Chem. 258,5707-5709. Kronman, M. J., and Bratcher, S. C. (1984).J.Biol. Chem. 259, 10887-10895. Kronman, M. J., and Holmes, L. G. (1965).Biochemistry 4,526-532. Kronman, M. J., Andreotti, R., and Vitols, R. (1964).Biochemistry 3, 1152- 1160. Kronman, M. J., Cerankowski, L., and Holmes, L. G. (1965).Biochemistry 4,518-525. Kronman, M. J., Holmes, L. G., and Robbins, F. M. (1967).Bzochzm. Biophys. Acta 133,

46-55.

Kronman, M. J., Holmes, L. G., and Robbins, F. M. (1971).J.Biol. Chem. 246, 1909-1921. Kronman, M. J., Hoffman, W. B., Jeroszko, J., and Sage, G. W. (1972a).Biochzm. Biophys. Acta 285, 124-144. Kronman, M. J., Jeroszko, J., and Sage, G. W. (197213).Biochim. Biophys.Acta 285, 145-166. Kronman, M. J., Sinha, S. K., and Brew, K. (1981).J. Biol. Chem. 256,8582-8587. Kumagai, I., Tamaki, E., Kakinuma, S., and Miura, K. (1987).J. Biochem. (Tokyo) 101,

51 1-517. Kurachi, K., Sieker, L. C., and Jensen, L. H. (1975).J. Biol. Chem. 250,7663-7667. Kuwajima, K. (1977).J. Mol. Biol. 114,241-258. Kuwajima, K. (1989).Proteins: Struct., Funct., and Genet. 6, 87- 103. Kuwajima, K., Nitta, K., Yoneyama, M., and Sugai, S. (1976).J. Mol. Biol. 106,359-373. Kuwajima, K., Hiraoka, Y., Ikeguchi, M., and Sugai, S. (1985).Biochemistry 24, 874-881. Kuwajima, K.,Harushima, Y., and Sugai, S. (1986).Int. J . Pep. Protein Res. 27, 18-27. Laird, J. E., Jack, L., Hall, L., Boulton, A. P., Parker, D., and Craig, R. K. (1988).Biochem.

J. 254,85-94.

La Rue, J. N., and Speck, J. C. (1970).J.Biol. Chem. 245, 1985-1991. Laschtschenko, P. (1909).Z. Hyg. Infektionskr. 64,419-426. Lavoie, T. B., Kam-Morgan, L. N. W., Hartman, A. B., Mallett, C. P., Sheriff, S., Saroff, D. A., Mainhart, C. R., Hamel, P. A., Kirsch,J. F., Wilson, A. C., and Smith-Gill, S. J.

LYSOZYME AND (Y-LACTALBUMIN

309

(1989a). I n “The Immune Response to Structurally Defined Proteins: The Lysozyme Model” (S. Smith-Gill and E. Sercarz, eds.), pp. 151-168. Adenine Press, Schenectady, New York. Lavoie, T. B., Kam-Morgan, L. N. W., Mallett, C. P., Schilling,J. W., Prager, E. M., Wilson, A. C., and Smith-Gill, S. J. (1989b). I n “The Use of X-Ray Crystallography in the Design of Anti-Viral Agents” (G. W. Laver and G. Air, eds.). Academic Press, San Diego, California. Lee, B., and Richards, F. M. (1971).J. M o l . B i d . 55,379-400. Lehmann, M. S., Mason, S. A., and McIntyre, G. J. (1985). Biochemistry 24, 5862-5869. Lewis, P. N., and Scheraga, H. A. (1971). Arch. Biochem. Biophys. 144,584-588. Lie, O., and Syed, M. (1986). Anim. Genet. 17,47-59. Lie, O., Solbu, H., and Syed, M. (1986). Anim. Genet. 17, 39-45. Lin, T.-Y. (1970). Biochemistry 9,984-995. Lindahl, L., and Vogel, H. J. (1984). Anal. Biochem. 140,394-402. LinderstrGm-Lang, K. (1952). “Lane Medical Lectures,” p. 53. Stanford Univ. Press, Stanford, California. Linse, S., Brodin, P., Johansson, C., Thulin, E., Grundstrom, T., and ForsCn, S. (1988). Nature (London) 335,65 1-652. Lonnerdahl, B., and Glazier, C. (1985).J.Nutr. 115, 1209-1216. Lord, R. C., and Yu, N.-T. (1970).J.Mol. Biol. 50,509-524. Luntz, T. L., Schejter, A., Garber, E. A. E., and Margoliash, E. (1989). Proc. Natl. Acad. Scz. U.S.A. 86,3524-3528. MacGillivray, R. T. A., Brew, K., and Barnes, K. (1979). Arch. Biochem. Biophys. 197, 404-4 14. MacKay, B. J., Iacono, V. J., Zuckerman, J. M., Osserman, E. F., and Pollock, J. J. (1982). Eur.J. Biochem. 129,93-98. Maes, E. D., Dolmans, M., Vincentelli,J. B., and Lkonis,J. (1969). Arch. Int. Physiol. Biochem. 77,388-390. Magee, S. C., and Ebner, K. E. (1974).J . Biol. Chem. 249,6992-6998. Magee, S. C., Mawal, R., and Ebner, K. E. (1973).J. Biol. Chem. 248,7565-7569. Magee, S. C., Geren, C. R., and Ebner, K. E. (1976). Bzochim. Biophys. Acta 420, 187-194. Maksimov, V. I., Kaverzneva, E. D., and Kravchenko, N. A. (1965).Biokhimiya (Moscou) 30, 866-872. Mann, G. (1906).“Chemistry of the Proteids,” pp. 360-361. Macmillan, London. Margoliash, E. (1963). Proc. Natl. Acad. Sci. U.S.A. 50,672-679. Matsumura, M., Becktel, W. J., and Matthews, B. W. (1988).Nature (London) 334,406-410. Matthews, B. W., Grutter, M. G., Anderson, W. F., and Remington, S. J. (1981). Nature (London) 290,334-335. Maurois, A. (1959). “The Life of Sir Alexander Fleming” (G. Hopkins, trans.), Cape, London, and Dutton, New York. McDonald, C. C., and Phillips, W. D. (1969). Biochem. Bzophys. Res. Commun. 35,43-51. McGuire, W. L. (1969). Anal. Biochem. 31,391-398. McKenzie, H. A. (1967). A h . Protein Chem. 22,55-234. McKenzie, H. A. (1970). In “Milk Proteins” (H. A. McKenzie, ed.), Vol. I, pp. 399-451. Academic Press, New York. McKenzie, H. A. (1971). In “Milk Proteins” (H. A. McKenzie, ed.), Vol. 11, pp. 474-476. Academic Press, New York. McKenzie, H. A. (1991). Submitted. McKenzie, H. A., and Ralston, G. B. (1971). Expetientia 27, 617-624. McKenzie, H. A., and Shaw, D. C. (1982). Proc. Int. C o n p . Biochem., 12th p. 325.

310

HUGH A. MCKENZIE A N D FREDERICK H . WHITE, JR.

McKenzie, H. A., and Shaw, D. C. (1985). Biochem. Znt. 10,23-31. McKenzie, H. A., and White, F. H., Jr. (1986). Anal. Biochem. 157, 367-374. McKenzie, H. A., and White, F. H., Jr. (1987). Biochem. Znt. 14,347-356. McKenzie, H. A., Muller, V. L., and Treacy, G. B. (1983). Comp. Biochem. Physiol. 74B, 259-27 1. McLean, D. M., Graham, E. R. B., Ponzoni, R. W., and McKenzie, H. A. (1984).J. Dairy Res. 51, 531-546. Messer, M., and Green, B. (1979). Awt. J. Biol. Sci. 32,519-531. Messer, M., and Kerry, K. R. (1973). Science 180,201-203. Messer, M., and Mossop, G. S. (1977). A w t . J . B i d . Sci. 30,379-388. Messer, M., Gadiel, P. A., Ralston, G. B., and Griffiths, M. (1983). Awt. J . Biol. Sn'. 36, 129-137. Metschnikoff, E. (1883). B i d . Zentralbl. 3,560-565. Meyer, K., Thompson, R., Palmer, J. W., and Khorazo, D. (1936). J . B i d . Chem. 113, 303-309. Miller, J. N., and King, L. A. (1975). Biochim. Biophys. Actu 393,435-445. Mitani, M., Harushima, Y., Kuwajima, K., Iweguchi, M., and Sugai, S. (1986).J.Biol. Chem. 261,8824-8829. Mitranic, M. M., and Moscarello, M. A. (1980). Cun.J. Biochem. 58, 809-814. Mitranic, M. M., and Moscarello, M. A. (1983). Prep. B i o c h . 13, 361-370. MorrC, D. J., Merlin, L. M., and Keenan, T. W. (1969). Biochem. Biophys. Res. Commun. 37, 813-819. Morrison, J. F., and Ebner, K. E. (1971a).J. Biol. Chem. 246,3977-3984. Morrison, J. F., and Ebner, K. E. (1971b).J. Biol. Chem. 246,3985-3991. Morrison, J. F., and Ebner, K. E. (1971c).J. B i d . Chem. 246,3992-3998. Morsky, P. (1988). Clin. Chim. Actu 178,327-336. Moser, l . , Pittner, F., and Dworsky, P. (1988).J . Biochem. Bqbhys. yethods 17,249-252. Moult, J., Yonath, A,, Traub, W., Smilansky, A., Podjarny, A., Rabinovich, D., and Saya, A. (1976).J . Mol. Biol. 100, 179- 195. Murakami, K., and Berliner, L. J. (1983). Biochemistry 22,3370-3374. Murakami, K., Andree, P. J., and Berliner, L. J. (1982). Biochemistry 21,5488-5494. Muraki, M., Morikawa, M., Jigami, Y., and Tanaka, H. (1987). Biochim.Biophys. Actu 916, 66-75. Musci, G., and Berliner, L. J. (1985a). Biochemistry 24,3852-3856. Musci, G., and Berliner, L. J. (198513).Biochemistry 24,6945-6948. Musci, G., and Berliner, L. J. (1986). Biochemistry 25,4887-4891. Musci, G., Koga, K., and Berliner, L. J. (1987). Biochemistry 27, 1260- 1265. Nagamatsu, Y., and Oka, T. (1980). Biochem.J. 185,227-237. Nagasawa, T., Kiyosawa, I., Fukuwatari, Y., Kitayama, T., Uechi, M., and Hyodo, Y. (1973). J . Dairy Sci. 56, 177- 180. Nicholas, K. R., Hartmann, P. E., and McDonald, B. L. (1981). Biochem.J. 194, 149-154. Nicholas, K. R., Messer, M., Elliott, C., Maher, F., and Shaw, D. C. (1987). Bi0chem.J. 241, 899-904. Nicholas, K., Loughnan, M., Messer, M., Munks, S., Griffiths, M., and Shaw, D. (1989). Comp. Biochem. Physiol. 94B, 775-778. Nicola, N. A., and Leach, S. J. (1976). Znt.J. Pept. Protein Res. 8,393-415. Nicolle, M. (1907). Ann. Znst. Pasteur 21,613-621. Nitta, K., and Sugai, S. (1989). Eur.1. Biochem. 182, 1 1 1 - 118. Nitta, K., Sugai, S., Prasad, R. V., and Ebner, K. E. (1984). Biochim. Biophys. Actu 786, 57-61.

LYSOZYME AND ff-LACTALBUMIN

31 1

Nitta, K., Tsuge, H., Sugai, S., and Shimazaki, K. (1987). FEBS Lett. 223,405-408. Nitta, K., Tsuge, H., Shimazaki, K., and Sugai, S. (1988). Biol. Chem. Hoppe-Seyler 369, 67 1-675. OKeeffe, E. T., Hill, R. L., and Bell, J. E. (1980a). Biochemistry 19,4954-4962. O’Keeffe, E. T., Mordick, T., and Bell, J. E. (1980b). Biochemistry 19,4962-4966. Ono, M., and Oka, T. (1980). Cell 19,473-480. Osserman, E. F., and Lawlor, D. P. (1966). J. Exp. Med. 124, 92 1-95 1. Osserman, E. F., Canfield, R. E., and Beychok, S. (eds.) (1974). “Lysozyme.” Academic Press, New York. Ostrovsky, A. V., Kalinichenko, L. P., Emelyanenko, V. I., Klimanov, A. V., and Permyakov, E. A. (1988). BiOphyS. C h . 30, 105-112. Palmer, K. J., Ballantyne, M., and Galvin, J. A. (1948).J . Am. Chem. Soc. 70,906-908. Pedersen, K. 0. (1936a). B z o c h . J . 30,948-960. Pedersen, K. 0. (193613).BzochmJ. 30,961 -970. Perkins, H. R. (1960). BiochemJ. 74, 182-186. Perkins, S. J., Johnson, L. N., Machin, P. A,, and Phillips, D. C. (1979). Biochem. J. 181, 2 1-36. Perkins, S. J., Johnson, L. N., Phillips, D. C., and Dwek, R. A. (1981). Biochem. J . 193, 553-572. Permyakov, E. A., Yarmolenko, V. V., Kalinichenko, L. P., Morozova, L. A,, and Burstein, E. A. (1981). Biochem. Biophys. Res. Commun. 100, 191-197. Permyakov, E. A., Morozova, L. A., and Burstein, E. A. (1985). Biophys. Chem. 21,21-31. Perrnyakov, E. A,, Murakami, K., and Berliner, L. J. (1987).J.Biol. C h a . 262,3196-3198. Pessen, H., Kumosinski, T. F., and Timasheff, S. N. (1971). J. Agnc. Food Chem. 19, 698- 702. Peters, C. W. B., Kruse, U., Pollwein, R., Grzeschik, K.-H., and Sippel, A. E. (1989). Eur. J . Biochem. 182, 507-516. Pfeil, W. (1981). Bzophys. Chem. 13, 181-186. Phillips, D. C. (1966). Sci. Am. 215,78-90. Phillips, D. C. (1967). Proc. Natl. Acad. Sci. U.S.A. 57,484-495. Phillips, D. C. (1974). In “Lysozyme” (E. F. Osserman, R. E. Canfield, and S. Beychok, eds.), pp. 9-30. Academic Press, New York. Phillips, D. C., Acharya, K. R.,Handoll, H. H. G., and Stuart, D. I. (1987). Bzochem. SOC. Trans. 15, 737-744. Phillips, N. I., and Jenness, R. (1971). Biochim. Biophys. Acta 229,407-410. Pierce, M., Turley, E. A,, and Roth, S. (1980). Znt. Rev. Cytol. 65, 1-47. Pincus, M. R., and Scheraga, H. A. (1981). Biochemistry 20,3960-3969. Podolsky, D. K., and Weiser, M. M. (1975). Biochem. Biophys. Res. Commun. 65, 545-551. Podolsky, D. K., McPhee, M. S., Alpert, E., Warshaw, A. L., and Isselbacher, K. J. (1981). N . Eng1.J. Med. 304, 1313-1318. Post, C. B., and Karplus, M. (1986).J. Am. Chem. SOC.108, 1317-1319. Post, C. B., Brooks, B. R., Karplus, M., Dobson, C. M., Artymiuk, P. J., Cheetham, J. C., and Phillips, D. C. (1986). J. Mol. Biol. 190,455-479. Poulsen, F. M., Hoch, J. C., and Dobson, C. M. (1980). Biochemistry 19,2597-2607. Powell, J. T., and Brew, K. (1974a). Biochem. J. 142,203-209. Powell, J. T., and Brew, K. (1974b). Eur.J. Bzochem. 48, 217-228. Powell, J. T., and Brew, K. (1976a). J. Biol. C h a . 951,3645-3652. Powell, J. T., and Brew, K. (1976b).J. Biol. Chem. 251, 3653-3663. Powell, J. T., and Brew, K. (1976~).Biochemistry 15,3499-3505. Powning, R. F., and Davidson, W. J. (1976). Comp. Biochem. Physiol. B . 55B, 221-228.

312

HUGH A. MCKENZIE AND FREDERICK H. WHITE, JR.

Prager, E. M., and Wilson, A. C. (1972). Biochem. Genet. 7,269-272. Prager, E. M., and Wilson, A. C. (1988).J. Mol. Evol. 27,326-335. Prager, E. M., Arnheim, N., Mross, G. A,, and Wilson, A. C. (1972). J . B i d . C h a . 247, 2905-2916. Prager, E. M., Wilson, A. C., and Arnheim, N. (1974).J. Biol. Chem. 249,7295-7297. Prasad, A. L. W., and Litwack, G. (1963). Anal. Biochem. 6,328-334. Prasad, R., and Ebner, K. E. (1980).J. B i d . C h a . 255,5834-5837. Prasad, R. V., Butkowski, R. J., Hamilton, J. W., and Ebner, K. E. (1982). Biochemistry 21, 1479-1482. Prieels, J.-P., and Schlusselberg, J. (1977). Biochim. Biophys. Acta 491, 76-81. Prieels, J.-P., Maes, E., Dolmans, M., and Leonis, J. (1975). Eur.J. Biochem. 60, 525-531. Prieels, J.-P., Dolmans, M., Schindler, M., and Sharon, N. (1976). Eur. J . Biochem. 66, 579-582. Prieur, D. J., and Froseth, L. G. (1986). Experientia 42,542-543. Proctor, V. A,, and Cunningham, F. E. (1988). CRC Crit. Rev. Food Sci. Nutr. 26,359-395. Proctor, S . D., Wheelcock, J. V., and Davies, D. T. (1974). B i o c h . Soc. Trans. 2,621-622. Ptitsyn, 0. B., Dolgikh, D. A., Gil’manshin, R. I., Shakhnovich, E. I., and Finkel’shtein, A. E. (1983). Mol. B i d . (MOSCOW) 17,569-576. Qasba, P. K., and Chakrabartty, P. K. (1978). J. Biol. Chem. 253, 1167- 1173. Qasba, P. K., and Safaya, S. K. (1984). Nature (London) 308,377-380. Qasba, P. K., Hewlett, I. K., and Byers, S. (1983). Biochem. Biophys. Res. Comm. 117, 306312. Quarfoth, G. J., and Jenness, R. (1975). Biochim. Biophys. Acta 379,476-487. Ram, B. P., and Munjal, D. D. (1985). CRC Crit. Rev. Biochem. 17,257-311. Rao, K. R., and Brew, K. (1989). B i o c h a . Biophys. Res. Comm. 163,1390-1396. Rawitch, A. B. (1972). Arch. Biochem. Biophys. 151,22-27. Rees, A. R., and Offord, R. E. (1972). B i o c h . J . 130,965-968. Remington, S. J., Anderson, W. F., Owen, J., Ten Eyck, L. F., Grainger, C. T., and Matthews, B. W. (1978).J. Mol. B i d . 118,81-98. Ries-Kautt, M. M., and Ducruix, A. F. (1989).J. Biol. Chem. 264, 745-748. Robbins, F. M., and Holmes, L. G. (1970). Biochim. Biophys. Acta 221, 234-240. Robbins, F. M., Kronman, M. J., and Andreotti, R. E. (1965). Biochim. Biophys. Acta 109, 223-233. Robbins, F. M., Andreotti, R. E., Holmes, L. G., and Kronman, M. J. (1967). Biochim. Biophys. Acta 133,33-45. Rodriguez, R., Menkndez-Arias, L., d e Buitrago, G. G., and Gavilanes, J. G. (1985). Biochem. Int. 11,841-843. Rodriguez, R., Menkndez-Arias, L., d e Buitrago, G. G., and Gavilanes, J. G. (1987). Comp. Biochem. Physiol. 88B,791-796. Rosenberg, S., and Kirsch, J. F. (1981). Biochemistry 20, 3196-3204. Rupley, J. A. (1967). Proc. R. SOC.London, Ser. B . 167,416-428. Sakar, P. K., Dutta, J., Sen, A., Phillips, N. I., and Jenness, R. (1971). J. Dairy Sci. 54, 120-122. Salton, M. R. J. (1952). Nature (London) 170,746-747. Salton, M. R. J. (1964). “The Bacterial Cell Wall,” Elsevier, Amsterdam. Salton, M. R. J., and Ghuysen, J. M. (1959). Biochim. Biophys. Acta 36,552-554. Salton, M. R. J., and Ghuysen, J. M. (1960). Biochim. Bqbhys. Acta 45,355-363. Sarma, R., and Bott, R. (1977).J . Mol. Biol. 113,555-565. Schachter, H., Jabbal, I., Hudgin, R. L., Pinteric, L., McGuire, E. J., and Roseman, S. (1970).J . Biol. Chem. 245, 1090- 1100.

LYSOZYME A N D ff-LACTALBUMIN

313

Schaer, J.-J., Milos, M., and Cox, J. A. (1985). FEBSLett. 190, 77-80. Schanbacher, F. L., and Ebner, K. E. (1970).J. Biol. Chem. 245,5057-5061. Scheraga, H. A. (1988). In “Biological and Artificial Intelligence Systems” (E. Clement and S. Chin, eds.), pp. 1-15. ESCOM, Leiden. Schindler, M., Sharon, N., and Prieels, J.-P. (1976). Biochem. Bzophys. Res. Commun. 69, 167- 173. Schmidt, D. V., and Ebner, K. E. (1971). Biochim. Biophys. Acta 243,273-283. Schmidt, D. V., and Ebner, K. E. (1972). Biochim. Biophys. Acta 268,714-720. Sebelien, J. (1885). 2. Physiol. Chem. 9,445-464. Secemski, I. I., and Lienhard, G. E. (1974).J. Biol. Chem. 249,2932-2938. Segawa, T., and Sugai, S. (1983).J. Biochem. (Tokyo) 93, 1321-1328. Selsted, M. E., and Martinez, R. J. (1980). Anal. Biochem. 109,67-70. Shakhnovich, E. I., and Finkelstein, A. V. (1982). Dokl. Akad. Nauk SSSR 267, 1247-1250. Sharma. R. N., and Bigelow, C. C. (1974).J. Mol. Biol. 88,247-257. Sheriff, S., Silverton, E. W., Padlan, E. A., Cohen, G. H., Smith-Gill, S. J., Finzel, B. C. and Davies, D. R. (1987). Proc. Natl. Acad. Sci. U.S.A. 84, 8075-8079. Sheriff, S., Silverton, E. W., Padlan, E. A., Cohen, G. H., Smith-Gill, S. J., Finzel, B. C., and Davies, D. R. (1988). In “Structure and Expression: From Proteins to Ribosomes” (R. H. Sarma and M. H. Sarma, eds.), Vol. 2, pp. 49-53. Adenine Press, Schenectady, New York. Shewale, J. G., Sinha, S. K., and Brew, K. (1984).J. Biol. Chem. 259,4947-4956. Shrake, A., and Rupley, J. A. (1980).Biochemistry 19,4044-4051. Shugar, D. (1952). Biochim. Biophys. Acta 8, 302-309. Silvia, J. D., and Ebner, K. (198O).J. Biol. Chem. 255, 11262-11267. Simons, E. R. (1981). In “Spectroscopy in Biochemistry” (J. E. Bell, ed.), Vol. I, pp. 63153. CRC Press, Boca Raton, Florida. Simpson, R. J., and Morgan, F. J. (1983). Biochim. Biophys. Acta 744,349-351. Simpson, R. J., Begg, G. S.,Dorow, D. S., and Morgan, F. J. (1980). Biochemistry 19, 1814- 1819. Sjogren, B., and Svedberg, T. (1930).J . Am. Chem. SOC.52,3650-3654. Smith, G. N., and Stocker, C. (1949). Arch. Biochem. 21,383-394. Smith, E. L., Kimmel, J. R., Brown, D. M., and Thompson, E. 0. P. (1955).J . Biol. Chem. 215,67-89. Smith, S. G., Lewis, M., Aschaffenburg, R., Fenna, R. E., Wilson, I. A., Sundaralingam, M., Stuart, D. I., and Phillips, D. C. (1987). Biochem. J. 242,353-360. Smith-Gill, S. J., Wilson, A. C.,Potter, M., Prager, E. M., Feldman, R. J., and Mainhart, C . R. (1982).J. Immunol. 128,314-322. Smith-Gill, S. J., Rupley, J. A., Pincus, M. R., Carty, R. P., and Scheraga, H. A. (1984). Biochemistry 23,993-997. Smolelis, A. N., and Hartsell, S. C. (1949).J. Bacteriol. 58, 731-736. Sommers, P. B., and Kronman, M. J. (1980).Biochim. Biophys. Acta 626,366-375. Sommers, P. B., Kronman, M. J., and Brew, K. (1973). Biochem. Biophys. Res. Commun. 52, 98-105. Sophianopoulos, A. J., and Van Holde, K. E. (1961).J. Biol. Chem. 236, PC82-PC83. Sophianopoulos, A. J., and Van Holde, K. E. (1964).J. B i d . Chem. 239,2516-2524. Sorensen, M., and Sorensen, S. P. L. (1939). C. R. Trav. Lab. Carlsberg, Ser. Cham. 23,55-99. Steinrauf, L. K. (1959). Acta Crystallop. 12,77-79. Stewart, C.-B., and Wilson, A. C. (1987). Cold Spring Harbor Symp. Quant. Biol. 52,891 -899. Stewart, C.-B., Schilling, J. W., and Wilson, A. C. (1987). Nature (London) 330, 401-404. Stewart, C.-B., Schilling, J. W., and Wilson, A. C. (1988). Nature (London) 332, 787-788.

3 14

HUGH A. MCKENZIE A N D FREDERICK H . WHITE, JR.

Strosberg, A. D., Nihoul-Deconinck, C., and Kanarek, L. (1970). Nature (London) 227, 1241-1242. Stuart, D. I,, Acharya, K. R., Walker, N. P. C., Smith, S. G., Lewis, M., and Phillips, D. C. (1986). Nature (London) 324,84-87. Sugai, S., Nitta, K., and Tsuge, H. (1988). In “Colloque (Institut National d e la Sante et la Recherche Medicale) 174, Deuxieme forum peptides” (A. Aubry, M. Maurraud, and B. Vitoux, eds.), pp. 591-596. John Libbey and Co., London. Takesada, H., Nakanishi, M., and Tsuboi, M. (1973).J.Mol. Biol. 77,605-614. Tamburro, A. M., Jori, G., Vidali, G., Scatturin, A., and Saccomani, G. (1972). Bzochim. Biophys. Acta 263,704-713. Tanahashi, N., Brodbeck, U., and Ebner, K. E. (1968). Biochim. Biophys. Acta 154,247-249. Tanford, C. (1970). A d z Protein C h a . 24, 1-95. Taniyama, Y., Yamamoto, Y., Nakao, M., Kikuchi, M., and Ikehara, M. (1988). Biochem. Biophys. Res. C m m u n . 152,962-967. Teahan, C. G. (1986). Ph.D. thesis. Aust. Natl. Univ., Canberra, Australia. Teahan, C. G., McKenzie, H. A., and Griffiths, M. (1991a). C m p . Bzochem. Physiol. I n press. Teahan, C. G., McKenzie, H. A., Shaw, D. C., and Griffiths, M. (1991b). Submitted. Teichberg, V. I., Kay, C. M., and Sharon, N. (1970). Eur.J. B i o c h . 16, 55-59. Teichberg, V. I., Sharon, M., Moult, J., Smilansky, A,, and Yonath, A. (1974).J. Mol. Biol. 87,357-368. Thompson, K., Harris, M., Benjamini, E., Mitchell, G., and Noble, M. (1972). Nature (London), New Biol. 238,20-2 1 . Thomsen, J., Lund, E. H., Kristiansen, K., Brunfeldt, K., and Malmquist, J. (1972). FEBS Lett. 22, 34-36. Trayer, I. P., and Hill, R. L. (1971).J.Biol. Chem. 246,6666-6675. Trayer, I. P., Barker, R., and Hill, R. L. (1974).I n “Methods in Enzymology” (W. B. Jakoby and M. Wilchek, eds.), Vol. 34, pp. 359-360. Academic Press, New York. Tsugita, A., and Inouye, M. (1968).J.Mol. Biol. 37, 201-212. Turkington, R. W., Brew, K., Vanaman, T. C., and Hill, R. L. (1968).J . Biol. Chem. 243, 3382-3387. Vanaman, T. C., Brew, K., and Hill, R. L. (1970).J.Biol. Chem. 245,4583-4590. Van Ceunebroeck, J.-C., Hanssens, I., Joniau, M., and Van Cauwelaert, F. (1985).J. Biol. Chem. 260, 10944- 10947. Van Ceunebroeck, J.-C., Krebs, J., Hanssens, I., and Van Cauwelaert, F. (1986).Biochem. Biophys. Res. Commun. 138, 604-610. Van Dael, H., Lafaut, J. P., and Van Cauwelaert, F. (1987). Eur. Biophys. J . 14, 409-414. Vernon, C. A. (1967). Proc. R . Soc. London, Ser. B 167,389-401. Vilotte, J.-L., Soulier, S., Mercier, J.-C., Gaye, P., Hue-Delahaie, D., and Furet, J.-P. (1987). Biochimie 69,609-620. Warme, P. K., Momany, F. A., Rumball, S. V., Tuttle, R. W., and Scheraga, H. A. (1974). Biochemistry 13,768-781. Watkins, W. M., and Hassid, W. Z. (1962).J.Biol. Chem. 237, 1432- 1440. Weaver, L. H., Grutter, M. G., Remington, S. J., Gray, T. M., Isaacs, N. W., and Matthews, B. W. (1985).J.Mol. E d . 21,97-111. Weiser, M. M., and Wilson, J. R. (1987). CRC Crit. Rev. Clzn. Lab. Sci. 14, 189-239. Weisman, L. S., Krummel, B. M., and Wilson, A. C. (1986).J.Biol. Chem. 261,2309-2313. Whitaker, J. R. (1963). Anal. Chem. 35, 1950- 1953. Whittaker, R. G., Fisher, W. K., and Thompson, E. 0. P. (1978). A w t . Zool. 20,57-68. White, F. H., Jr. (1976). Biochemistry 15, 2906-2912. White, F. H., Jr. (1982). Biochemistry 21,967-977.

LYSOZYME AND ff-LACTALBUMIN

315

White, F. H., Jr., and Wright, A. G., Jr. (1984). Znt. J. P e p . Protein Res. 23, 256-270. White, F. H., Jr., McKenzie, H. A., Shaw, D. C., and Pearce, R. J. (1988). Biochem. Znt. 16, 521-528. White, T. J., Mross, G. A., Osserman, E. F., and Wilson, A. C. (1977). Biochemistry 16, 1430-1436. Wichmann, A. (1899).Z. Physiol. Chem. 27, 575-593. Wilmut, I., Clark, J., and Simons, P. (1988). New Sci. 119,56-59. Wilson, A. C., Carlson, S. S., and White, T. J. (1977). Annu. Rev. Biochem. 46, 573-639. Wilson, A. C., Ochman, H., and Prager, E. M. (1987). Trends Genet. 3,241-247. Wong, L.-J. C., and Wong, S. S. (1984). Znt.1. Biochem. 16, 913-917. Woods, K. L., Cove, D. H., and Howell, A. (1977). Lancet 2, 14-16. Yasunoba, K. T., and Wilcox, P. E. (1958).J.Biol. Chem. 231,309-313. Yu, N.-T. (1974).J.Am. Chem. SOC.96,4664-4668. Yu, N.-T. (1977). CRC Crit. Rev. Biochem. 4,229-280. Zeng, J., Rao, K. R.,Brew, K., and Fenna, R. (1990).J . Biol. Chem. 265, 14886- 14887. Zuckerkandl, E. (1987).J. Mol. Euol. 26,34-46. Zuckerkandl, E., and Pauling, L. (1962). In “Horizons in Biochemistry” (M. Kasha and B. Pullman, eds.), pp. 189-225. Academic Press, New York. Zuckerkandl, E., and Pauling, L. (1965). In “Evolving Genes and Proteins” (V. Bryson and H. J. Vogel, eds.), pp. 97- 166. Academic Press, New York. NOTEADDEDI N PROOF.Model 11, as discussed in Section X,B,4, was suggested first by Dayhoff (1972) prior to its development and extension by Wilson et al. (1977). T h e most recent refinement follows from the first determination of sequences of fish lysozymes (rainbow trout, Oncorhynchzcs mykiss), resulting in a new proposed divergence date of ca.400 MY ago (Daotigny et al., 1991). Daotigny, A., Prager, E. M., Pham-Dinh, D., Jollks, J., Pakdel, F., Grinde, B., and Jollt5s, P. (1991).J.Mol. E d . 82, 187-198. Dayhoff, M. O., ed. (1972). “Atlas of Protein Structure and Sequence,” Vol. 5, pp. D133-D-135. National Biomedical Research Foundation, Silver Spring, Maryland.

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AUTHOR INDEX

Numbers in italics refer to the pages on which the complete references are listed.

A Abdel-Meguid, S. S., 24, 35 Abraham, E. P., 176,299 Abraham, F. F., 132, 161 Acharya, K. R., 189, 193, 209,210,211, 212, 213, 223, 239, 247, 248,267,281, 283,288,293,299,306,311,313 Achter, E. K., 265,299 Acker, L., 94, 162, 169 Adair, G. S., 61, 162 Adair, M. E., 61, 162 Adebodun, F., 297,299 Adler, J., 156, 158, 159, 164 Adolph, G. R.,241,302 Ahlstroem, P., 97, 113, 114, 130, 162 Alber, T., 142, 162, 295,299 Alderman, E. M., 298,304 Alderton, G., 197,299 Alexander, S., 66, 67, 164 Ali, S., 54, I62 Aliotta, F., 110, I62 Allison, V. D., 176, 304 Almog, R.,46, 126, 127, 162 Alnemri, E. S., 296, 307 Alpert, E., 251,311 Althoff, S., 22,33 Amano, T., 23 1, 245, 246, 308 Amit, A. G., 274,299 Anagnostopoulou-Konsta, A., 68,69, 162 Anderson, W. F., 23, 24,33,283,309,312 Andersson, L., 118, I72 Andersson, T., 72, 76, 166 Andree, P. J., 217, 219, 254, 257, 262, 297, 299,310 Andreeva, A. P., 76,88, 136, 162 Andreotti, R.,267,308 Andreotti, R. E., 260, 267,312

Andrew, E.R.,71,73, 136, 162, I 6 5 Andrews, A. T., 252,299 Andrews, L. J., 260,306 Andrews, P., 251,253,255,299, 307 Andronikashivili, E. L., 50, 162 Anfinsen, C. B., 269, 277,300 Angelis, L. D., 112, 163 Aqvist, J., 143, I72 Arakawa, T., 5 , 6 , 3 3 , 36 Archer, M., 278,300 Argarana, C. E., 8, 35 Armstrong, J. M., 187, 300 Arnebrant, T., 59, 164 Arnheim, N., 181,231,244,273,300,311 Arnon, R., 273,300,301 Artymiuk, P., 195, 301 Artymiuk, P. J., 99, 100, 101, 102, 104, 112, 127, 144, 146, 163,169, 194,203, 204,294,300,311 Asai, M., 17, 33 Aschaffenburg, R., 189, 197, 209, 210, 300,313 Ashford, V., 2 1 , 3 4 Assaf, Y., 202, 312 Astier, J. P., 7, 8, 13,33 Ataka, M., 8, 17, 18, 26, 33, 68, 110, 162 Atanasov, B. P., 76, 88, 136, I62 Atassi, M. Z., 271,273,300 Aumann, K. D., 103,169 Aune, K. C., 263,300 Aviram, I., 110, 162

B Babad, H., 178,253,300 Bachleitner, A., 43,49, 89, 109, 110, 130, 131, 136, 145,164 317

318

AUTHOR INDEX

Bagshaw, W., 27 1,300 Baianu, 1. C., 75, 76, 167, 168 Baird, J. K., 21, 34 Bajaj, M., 178, 283, 300 Baker, E. N., 5,33,99, 104,162 Baker, L. J., 81, 84, 162 Baker, T., 104, 162 Balaram, P., 99, 167 Baldwin, E. T., 13, 25, 33 Ballantyne, M., 197,311 Ballardie, F. W., 183, 300 Banerjee, S. K., 16, 17,33, 144, 162, 201, 260,300,306 Banyard, S. H., 197,300 Barbaric, S., 95, I62 Barel, A. O., 208,264,300 Barford, D., 283,306 Barker, R., 251,300,314 Barlow, D. J., 99, 109, 162 Barman, T. E., 225,271,272,276,294,300 Barnes, K., 227,238,309 Bartunik, H., 103, 169 Bash, P. A,, 121, 162 Baum, J., 269,270,300 Bauminger, E. R., 88, 162 Bechtold, G., 62, 165 Becktel, W. J., 295, 309 Beece, D., 96, 130, 162 Beg, 0. U., 189,227,240,300 Begg, G. S., 181,231,300,313 Behi, J., 66, 162 Bell, J. E., 253,254,255,257,301,310 Bell, K., 189, 227,232,238,292,300,301, 305 Bell, R. P., 143, I62 Belonogova, 0. V., 76, 88, 136, 162 Ben-Naim, A,, 119, 120,162 Benjain, D. C., 273,274,278,301 Benjamini, E., 273,314 Bennich, H., 282-283,303 Bentley, G. A., 100, 168 Benton, M., 278,279,301 Berezin, 1. V., 96, 168 Bergenstaahl, B., 59, 164 Berger, L. R., 178,301 Berlin, E., 46, 54, 162 Berliner, L. J., 213,217,219,220,223, 251, 254, 257,262,266,297,299,301, 305,308,310,311 Bernhardt, J., 50, 162

Bernier, l., 231, 243, 244, 306 Bernstein, H., 263,305 Berthou, J., 195,301,306 Berzofsky,J. A., 273,274,278,301 Bettelheim, F. A., 54, 162 Betts, L., 18,26, 33 Betzel, C., 22,35, 105, 162 Beveridge, D. L., 120, 121, 168 Beychok, S., 175,263,305,310 Beyer, T. A., 254,257,301 Bhattacharjee, N., 189,301 Bickham, D., 25,34 Bigelow, C. C., 271,313 Bjorck, L., 182, 303 Blackburn, D. G., 292,305 Blake, C. C. F., 99, 100, 101, 102, 104, 127, 144, 146,163, 174, 192, 193, 194, 197, 198,203, 204, 212, 213, 244, 264, 266,294,300,301 Blanc, B., 52, 54, 170 Blevins, R. A., 105, 146, 163 Bloomfield, A., 176,301 Blout, E. R., 270, 271,304 Blow, D. M., 106, 163 Blumberg, B. S., 189,301 Blundell, T., 178, 283 Blundell, T. L., 2,5,20, 26,33, 104, 162, 195,301 Boesman-Finkelstein,M., 182, 301 Bohme, D. K., 143,163 Boistelle, R.,7, 8, 13, 33 Boman, H. G., 282-283,303 Bone, S., 64,66, 162, 163 Bonnemaire,J., 189,305 Boodhoo, A., 23,24,33 Borah, B., 74, 163 Bordet, M., 176,301 Bordet, M. J., 176,301 Bott, R.,202,312 Bott, R. R., 24, 33 Boudouris, G., 68, 167 Boulton, A. P., 281, 308 Bourret, D., 74,163 Bowman, B. H., 229,242,243,285,307 Bradbury, E. M., 266,303 Bradbury, J. H., 266, 27 1,301 Bragg, L., 174,301 Brasch, J. W., 109, 166 Bratcher, S. C., 217, 218, 219, 262,287, 301,308

319

AUTHOR INDEX

Brayer, G. D., 23,35 Brazhnikov, E. V., 276, 294,303 Breese, K., 41.49, 50, 54, 126, 127, 141, 163 Brennan, R. G., 20, 33 Breslow, E., 22, 35 Brew, K., 175, 179, 180, 189, 206, 208, 210,217,218,219, 223,227,232, 238, 239,240,248, 250, 251,252,253,254, 255,256,257, 261,262,263,266,269, 271,280,287, 288,290,292,297,301, 302,303,304,305,307,308,309, 311, 312,313,314,315 Brick, P., 24, 25,33,35, 106, 163 Bridelli, M.G., 68, 163 Brodbeck, U., 179, 191,255,270,302, 303,304,313 Brodin, P., 288, 309 Brooks, B., 87, 163 Brooks, B. R., 112, 169,203,311 Brooks, C. L., 112, 163 Brooks, C. L., 111,205,302 Broom, M. B., 26,29,33-34 Broom, M. B. H., 16,33 Brown, D. M., 181, 185,313 Brown, F., 121, 167 Brown, F. K., 121, 162 Brown, J. R., 244,280,302,305 Brown, K. A., 106, 163 Brown, R. C., 189,227,302 Brown, W. E., 110, 163 Browne, W. J., 206, 207,208,210, 255, 266,27 1,302 Bruccoleri, R. E., 87, 163 Bruice, T. C., 143, 163 Brunauer, S., 43, 163 Brunfeldt, K., 229, 241, 314 Bruni, F., 66, 67, 69, 150, 170 Brusca, A. E., 182,302 Bryan, W. P., 42,44,81,84,162,163,170 Bryant, D. J., 71,162,165 Bryant, R. G., 55,71, 72, 73, 74, 97, 128, 136, 163,166, 169,171 Bugg, C. E., 26,29, 32,33, 33-34,34 Bull, H. B., 41, 49, 50, 54, 126, 127, 141, 163 Burcham, T. S., 151, 165 Burditt, L. J., 299, 302 Burstein, F. A., 85, 169, 216, 217, 221, 261,297,311

Buss, D. H., 197,209,300 Butkowski, R. J., 189, 227,240, 311 Bychkova, V. E., 276,294,303

C Calvert, P. D., 13, 14, 16, 17, 34 Cameron, I. L., 74,75, 165 Campbell, K. S. W., 276, 302 Campbell, P. N., 175, 227, 239, 281, 282, 298,299,302,303,305 Canfield, R., 273,300 Canfield, R. E., 175, 181, 183, 201, 229, 231,240-241,243,244,302,310 Cantor, C. E., 8 , 3 5 Capelletti, R., 68, I63 Capon, B., 183,300 Careri, G., 39, 44, 49, 50, 64, 65, 66, 67, 69, 81, 85, 93, 107, 108, 109, 112, 122, 125, 130, 131, 135, 141, 145, 149, 150, 163, 170,263,302 Carlson, S. S., 277, 286, 314 Carlson, W. D., 105, 144, 171 Carr, C. W., 215,302 Carroll, R. L., 278,302 Carter, C. W., Jr., 2, 13, 18, 25, 26,33 Carter, D., 26,29,33-34 Carter, D. B., 8, 33 Carter, D. C., 16, 20, 29, 33, 34 Carter, J. B., 8, 33 Carty, R. P., 202, 313 Cashell, E. M., 71, 165 Caspar, D. L. D., 5,33, 104, 163 Castan6n, M. J., 24 1, 302 Castellino, F. J., 180, 191, 208, 302 Cavarelli, J., 25, 35 Cavatorta, F., 150, 163 Celaschi, S., 68, 163 Cerankowski, L., 187, 259, 260, 261, 308 Cerofolini, G. F., 44, I63 Cerofolini, M., 44, 163 Chain, E., 177,303 Chakrabartty, A., 151, 172 Chakrabartty, P. K., 189,312 Chandan, R. C., 183,302 Chandler, D. K., 253,302 Chase, J. W., 24,35 Chau, K. H., 264,302 Cheetham, J., 283,306

320

AUTHOR INDEX

Cheetham, J. C., 203,311 Chen, M. C., 264.302 Chen, Y.-H.. 264, 302 Chiarandra, G., 271,302 Chin, S., 41, 164 Chipman, D. M., 202,312 Chirgadze, Y. N., 80, 97, 111, 131, 140, 148,163,164 Chlebowicz-Sledziewska, E., 241, 302 Chou, P. Y., 273,302 Chung, L. P., 241,302 Churg, A. K., 122,164, 171 Claesson, P. M., 59, 164 Clancy, L. L., 8, 33 Clarage, J., 5, 33, 104, 163 Clarage, M., 5,33, 104, 163 Clark, J., 298, 314 Clegg, J. S., 95, 151, 164 Clemens, W. A., 278,279,280,302 Clementi, E., 41, 120, 164 Clerc, J. P., 66, 67, 164 Clore, G. M., 267,305 Clymer, D. C., 253,302 Cohen, G. H., 274,312 Cole, A. G., 49, 166 Cole, G., 105, 144, 168 Collins, J. C., 183, 302 Colvin, B., 191, 255,304 Conti, A., 189, 222, 227, 229, 240, 241, 302,304 Cook, S. P., 295, 299 Corey, M. J., 203,307 Cornish-Bowden, A., 224,284,302,303 Cortopassi, G. A,, 185, 303 Cove, D. H., 250, 315 Cowburn, D. A., 263,266,303 Cox, J. A., 218, 312 Cox, M. J., 8, 26, 27,30,33,36 Craig, R. K., 227, 239,281,282, 298,299, 302,303,305,308 Crane-Robinson, C., 266,303 Creighton, S., 143, 172 Creighton, T. E., 192, 270,271,303,304 Crespi, H. L., 85,86, 168 Critchlow, J. E., 143, 162 Croizier, G., 181,282,307 Croizier, L., 181, 282,307 Crompton, A. W., 278,303 Cross, M., 241, 281,303 Crowe, J. H., 95, 151, 164

Crumley, K. V., 13, 33 Cullis, P. M., 118, 172 Cummingham, F. E., 175,312 Cunningham, W. P., 250,303 Curry, K. A., 8, 33 Cusack, S., 85, 86, 87, 130, 136, 164, 171 Cutfield, J. F., 104, 162 Cutfield, S. M., 104, 162 Cygler, M., 24, 33

D Daggett, V., 121, 167 Dandekar, A. M., 240,281,303 Dao-pin, S., 204,295,299,303 Darby, G., 29,34 D’Arcy, R. L., 4 1-42, I72 Das, B., 44, 170 Dauter, Z., 22,35 Davey, R. J., 13,33 Davidson, W. J., 181,311 Davies, D. R., 25, 34, 105, 144, 171, 274, 313 Davies, D. T., 189, 190, 312 Davies, M. S., 281, 298, 305 Davis, M. E., 257, 301 Dawson, T. J., 280, 303 Day, M. F., 276, 302 De Buitrago, G. G., 231,246, 288,312 De Gennes, P. G., 156,164 Decter, J. B., 3 1, 35 Deibel, M. R., 8, 33 Deininger, C. A., 152, 172 Delaney, R., 280, 305 Delbaere, L. T. J., 24, 35 Delepierre, M., 80, 164 DeLucas, L. J., 21, 26, 29, 32, 33, 33-34, 34

DeMattei, R. C., 13, 14, 16, 19,32,34,36 Denton, W. L., 179,302,303 Deonier, R. C., 267, 303 Desmet, J., 220, 223, 261, 303 Deutscher, G., 156, 158, 159, 164 Deutschmann, G., 80, 164 DeVries, A. L., 151, 164, 167, 170 Dhawale, S. W., 251, 305 Dianoux, A. C., 181,303 Dickerson, R. E., 194,287,303,307 Dickie, H. M., 197, 300

AUTHOR INDEX

Dickman, S. R., 185,303 Dijkstra, B. W., 2 2 , 2 6 , 3 4 , 3 5 Dissado, L. A., 68, 170 Dobson, C. M., 80, 112, 164, 169, 203, 213,216,266,269,270,300,303,311 Dobson, D. E., 185, 229,242, 284,303, 307 Dodson, E., 104, 162 Dodson, E. J., 104, 162 Dodson, G., 104, 162 Dodson, G. G., 104, 162 Dolgikh, D. A., 269, 276,294,303,312 Dolmans, M., 252,257,271,309,312 Dorland, L., 292,307 Dorow, D. S., 181,231,313 Doster, W., 43,49, 87, 89, 109, 110, 130, 131, 136, 145, 163, 171 Douzou, P., 198,303 Downer, N. W., 51, 81, 82,83, 140, 141, 170 Dransfeld, K., 62,64, 89, 130, 136, 171 Drapon, R., 94, 164 Dratky, O., 74, 164 Draut, J., 2 1, 34 Drenth, J., 22, 24, 26,34, 36,41, 168 Dreusicke, D., 5 , 3 4 Drewry, J., 189,200 Duckworth, R. B., 152, 164 Ducruix, A. F., 197,312 Dunau, R., 43,49,89, 109, 110, 130, 131, 136, 145,164 Durbin, S. D., 14, 16,34 Durchschlag, H., 74, 164 Dutta, J., 272, 312 Dwek, R.A., 199,216,267,307,311 Dworsky, P., 185,310

E Eadie, G. S., 185, 307 Ealick, S. E., 29, 34 East, I. J., 273, 274, 278, 301 Eaton, W. A,, 8, 12, 18, 34, 35 Ebner, K., 253,313 Ebner, K. E., 179, 180, 189, 191,227, 240, 252,253,255,256,257,272,301,302, 303,304,309,310,311,312,313 Edbrooke, M. R., 227,239,281,282,305 Eden, J., 66, 163, 164

32 1

Edsall, J. T., 38,40,41,99, 117, 118, 164, 194,204, 224,262,303,304 Edwards, S. L., 21,34 Ehrenberg, A,, 41,164 Ehrenpreis, S., 182,303 Eichele, G., 23, 36 Eigner, E. A., 147, 168 Einspahr, H. M., 8,33 Eisenberg, D., 41, 61, 118, 142, 164, 170 Eisenstadt, M., 76, 164 Eisenstein, L., 96, 130, 162 Eklund, H., 147, 171 Ekstrand, B., 182,303 Elliot, C., 189, 310 Ellis, P. D., 219, 266, 301 Emelyanenko, V. I., 262, 311 Emery, D. C., 281,305 Emmett, P. H., 43, 163 Engelmann, H., 90,130,167 Engstrom, A., 282-283,303 Epstein, C. A., 177, 303 Erdmann, V. A., 2 2 , 3 5 Essam, J. W., 156, 164 Evans, P. A,, 269,270,300

Fainanu, M., 273,304 Farris, J. S., 288, 304 Fasella, P., 130, 163 Fasman, G. C., 264, 305 Fasman, G. D., 273,302 Faure, A., 273, 288,304 Fedotov, V. D., 62, 165 Feeney, R. E., 151, 165 Feher, G., 7, 12, 13, 14, 16, 19, 32, 34, 35 Fehribach, J. D., 21, 32, 34 Feigelson, R. S., 2, 13, 14, 15, 16, 19, 32, 34,36 Feldman, R. J., 274,275,313 Fel'dman, Y. D., 62,165 Fellows, R. E., Jr., 280, 305 Fenna, R., 288,315 Fenna, R. E., 190, 197,209, 210,300,304, 313 Fenna, R. H., 174, 192,201 Fernandez-Sousa, J. M.,283,304 Fett, J. W., 298,304 Fevold, H. L., 197,299

322

AUTHOR INDEX

Fiddis, R. W., 13, 14, 16, 17, 34 Fiess, H. A,, 215, 304 Findlay, J. B. C., 189, 227, 239, 304 Findsen, E. W., 110, 165 Finkel’shtein, A. E., 269, 312 Finkelstein, A. V., 269, 313 Finkelstein, R. A., 182, 301 Finney, J., 85,86,87, 164, I71 Finney, J. L., 42, 55, 83, 99, 108, 109, 122, 165,169 Finzel, B. C., 5, 34, 104, 165, 274, 313 Fish, W. W., 189,227,302 Fisher, W. K., 292, 314 Fitch, W. M., 288-289,304 Fitzgerald, D. K., 191, 255, 304 Fitzgerald, P. M. D., 23, 34 Flannery, T. F., 278, 300 Fleming, A,, 176, 304 Fleming, G. R., 122, 168 Fletcher, P. D. I., 95-96, 165 Flewelting, R. F., 122, 166 Flory, P. J., 43, 154, 165 Fokker, A. P., 176,304 Fontana, M. P., 110, 150,162, 163 Fontecilla-Camps,J. C., 5, 34 Ford, L. O., 201,304 Forsen,S., 72, 76,97, 113, 114, 130, 162, I66 Forsen, S., 216, 288, 307, 309 Forster, L. S., 85, 111, 165, 171, 260, 306 Foss, J. G., 45, 165 Fouche, P. B., 181,304 Fowlis, W. W., 21,34 Franks, F., 41, I65 Fraser, 1. H., 252, 304 Frauenfelder, H., 96, 130, 148, 149, 162, I65 Freed, J. H., 142, 172 Freedman, R. B., 95-96, 165 Frey, M., 5, 34 Frick, L., 18.25, 26, 33 Friedberg, F., 213, 304 Frolov, E. N., 76, 88, 136, 162 Fromageot, C., 229,306 Froseth, L. G., 284,312 Fucaloro, A. F., 85, 165 Fujio, H., 231, 245, 246, 308 Fujita, Y., 52, 60, 140, I65 Fukuwatari, Y., 189, 310 Fuller, N., 58, 168

Fuller, N. L., 40,56,57,58, 129, 138, 169 Fullerton, G. D., 74, 75, 165 Funatsu, M., 260,305 Fung, B. M., 71,165 Furet, J.-P., 227, 232,239, 281,304, 314 Furuno, T., 2, 34

G Gabellieri, E., 84, I71 Gabriel, C., 62, I66 Gadiel, P. A., 292, 310 Galat, A., 270, 271, 304 Galiazzo, G., 215, 307 Gallo, A. A., 215, 266, 304 Calvin, J. A,, 197, 311 Gao, J., 121, I65 Garavito, R. M., 2, 34 Garber, E. A. E., 295, 309 Gardiner, B. G., 278,279,304 Garfinkel, D., 262,304 Garson, J. C., 68, 167 Gascoyne, P. R. C., 42,43,66, 164, I65 Gaspar, R., Jr., 71, 165 Gaubman, R. E., 90, 130, 167 Gauthier, J., 278,304 Gavilanes, J. G., 231, 246, 273, 283, 288, 304,312 Gavish, B., 61, 96, 130, 165 Gawinowicz Kolks, M. A., 8, 35 Gaye, P., 227,232, 239,281, 304, 314 Gayen, S. K., 197,300 Geige, R., 23, 35 Geiger, A., 112, 165 Geis, I., 287, 303 Gekko, K., 60, 167 Gelin, B. R., 98, 168 Gennari, G., 215, 307 Genovesio-Taverne, J.-C., 5, 34 Genzel, L., 62,64, 165, 167, 169 Geren, C. R., 252, 253,302,309 Gernert, K. M., 20,34 Getova, T., 299,302 Gevorkyan, S. G., 43,98, 136,165,168 Ghuysen, J. M., 177,312 Giannini, L., 110, 165 Giansanti, A., 49,50,64,65,66,67,69, 107, 108, 109, 131,135, 141, 145, I63 Giblett, E. R., 185, 304

323

AUTHOR INDEX

Gibson, K. D., 119, 167, 294,304 Giege, R.,21, 23, 25, 32, 34, 35 Gilliland, G. L., 8, 25, 34, 104, 172 Gil'manshin, R. I., 269, 304, 312 Gil'manshin, R. J., 276, 294, 303 Gilson, M. K., 122, 165 Gins, V. K., 76, 88, 136, 162 Ginzberg, B. Z.,,46, 166 Giordano, R.,110, 162 Giraud, G., 66,67, 164 Glaser, M., 22,33 Glazer, A. N., 185,263,304,306 Glazier, C., 223, 309 Godovac-Zimmermann, J., 189,222, 227, 229,240,241,302,304 Gol'danskii, V. I., 76, 88, 90, 130, 136, 162,167 Goldanskii, V. I., 88, 90, 91, 136, 137, 166, 167 Gonnelli, M., 96, 166, 171 Good, D., 96, 130,162 Goodfellow, J. M., 55, 99, 122, 165, 171 Gordon, S., 241,302 Gordon, W. G., 180, 189, 190,304 Grace, D. E. P., 294,300 Graeslund, A., 41, 164 Grainger, C. T., 283, 312 Grant, E. H., 62, I66 Gratner, W. B., 263,266,303 Gratton, E., 39,44, 49,50, 61,69, 81, 85, 93, 107, 108, 109, 110, 112, 122, 125, 130, 131, 141, 149, 163,165, 170,263, 302 Gray, T. M., 204, 283,314 Greenfiekl, N., 264,305 Gregory, R. B., 80, 147, 166, 168 Grez, M., 281,307 Griffiths, M., 189, 292,310 Griffiths, M. E., 182, 189, 222, 229, 243, 247,280,287,291,305,314 Grinde, B., 182,305 Groendijk, H., 24,36 Gronenborn, A. M., 267,305 Gros, P., 30, 35 Grosclaude, F., 189, 305 Griitter, M. G., 283,305,309 Grundstroom, T., 288,309 Grunwald, J., 25 1,305 Griitter, M. G., 204,283,314 Grzeschik, K.-H., 241, 281, 311

Guddat, L. W., 24, 35 Curd, F. R. N., 273,274,278,301 Guy, H. R., 80, 118, 166 Guyon, E., 66,67, 164

H Habeeb, A. F. S., 273,300 Haggis, G. H., 80, 166 Hagler,A. T., 100, 115, 116, 142, 166 Haire, R. N., 146, 166 Hall, L., 175, 227,239, 281, 282, 298, 299, 303,305,308 Halle, B., 72, 76, 166, 169 Hallenga, K., 72, 167 Halliday, J. A., 189, 305 Halloran, T. P., 21, 34 Halper, J. P., 263, 305 Haly, A. R., 50, 54, 166 Hamaguchi, K., 215,306 Hamano, M., 218,297,305 Hamel, P. A., 275, 308 Hamilton, D. W., 258,299,305 Hamilton, J. W., 189, 227, 240, 311 Hamm, R.,94,169 Hammer, M. F., 183,305 Handford, B. O., 197,209,300 Handoll, H. H. G., 193,209,213,311 Hanley, C., 269, 270,300 Hannum, C., 273,274,278,301 Hansen, A. M. F., 81, 84, 162 Hanson, J. C., 103, 166 Hanson, L. A,, 189,305 Hanssens, I., 217, 220,221, 261,271,303, 305,314 Hara, S., 283, 305 Harada, S., 283,305 Harburn, G., 20, 34 Hardman, K. D., 104,165 Hardy, C. J., 61,165 Harel, M., 106, 171 Harris, M., 273, 314 Harris, P. K., 8, 33 Harrison, P. M., 88, 162 Hartley, B. S., 280, 305 Hartman, A. B., 275,308 Hartmann, H., 103, 166, 169 Hartmann, P. E., 189,310 Hartsell, S. C., 185, 313

324

AUTHOR INDEX

Harushima, Y., 220,297,308,310 Harvey, S. C., 62, 71, 128, 166 Hash, J. H., 181,304 Hassid, W. Z., 178, 253, 300, 314 Hawkes, J. J., 64, 166 Hayashi, K., 260, 305 Hayashi, S., 17,36 Hayssen, V., 292,305 Hedlund, B. E., 146, 166 Heidemeier,J., 88, 89, 136, 169 Heidner, E., 18,26, 34 Heinrikson, R. L., 8, 33 Helliwell, J. R., 26, 34 Hendrickson, W. A.. 5,8,35,36, 102, 166 Hermann, J., 231,245,246,305 Hermans, J., 115, 117, 142,166 Herreman, W., 271,305 Herren, B., 26,29,33-34 Herries, D. G., 256, 257, 307 Hess, G. P., 144, 162, 201,300 Hew, C. L., 151, 172 Hiebl, M., 43, 49, 89, 109, 110, 130, 131, 136, 145, 164 Hill, R. L., 175, 179, 180, 189, 191, 206, 207,208, 210,227,232, 250, 251, 253, 254, 255,257,266,271,280,292,300, 301,302,305,310,314 Hill, T. L., 43,44, 49, 134, 166 Hiltner, A., 98, 166 Hilton, B. D., 71, 72, 80, 166, 172 Hindenburg, A,, 231,300 Hirai, Y., 8, 34 Hiraoka, Y., 221, 265, 269, 305, 306,308 Hirs, C. H. W., 2,36,41, 166 Hirsch, E., 23, 32, 34 Hirsch, R. E., 19, 3 4 Hnojewyj, W. S., 45, 80, 127, 166 Hoa, G. H. B., 198,303 Hoch, J. C., 213,266,311 Hodgkin, D., 104, 162 Hodgkin, D. M. C., 104, 162 Hoekstra, P., 62, 71, 128, 166 Hoffman, W. B., 208,218,219,260,308 Hofrichter, J., 8, 12, 18, 34, 35 Hol, W. G. J., 24, 30, 35, 36 Holden, H. M., 105, 166 Holladay, L. A,, 267, 306 Holler, E., 144, 162, 201, 300 Holmes, L. G., 187, 208, 259, 260, 261, 264,267,308,312

Honig, B. H., 122, 165, 166 Hope, H., 102,171 Hopp, T. P., 189, 227, 239, 273, 306 Hopper, K. E., 187, 189, 190, 191,222, 227,229,232,238,264,291,300,301, 306 Howard, J. B., 185,306 Howard, S. B., 21,34 Howarth, M. A., 80, 164 Howell, A., 250, 315 Hsi, E., 55, 71, 72, 74, 166 Hubbard, R., 104,162 Hubbard, R. E., 99, 104,162 Hubbell, W. L., 122, 166 Hudgin, R. L., 250,312 Hudson, B. G., 189,227,302 Hue-Delahaie, D., 227, 232, 239, 281, 304, 314 Hunklinger, S., 62, 64, 89, 130, 136, 171 Hurley, W. L., 281,306 Hutchens, J. O., 49, 166 Hvidt, A., 80, 166 Hwang, J. K., 121,172 Hymes, A. J., 191,306 Hyodo, Y., 189, 310

I Iacono, V. J., 182, 309 Ibrahimi, I. M., 231, 244,245,296,306, 307 Ikeda, K., 215,306 Ikeguchi, M., 265,269, 271,297,306,308, 310 Ikehara, M., 270,271, 314 Ikenaka, T., 283,305 Illyustrov, N. V., 88, 162 Imoto, T., 175, 193,201, 202,216,229, 244,260,305,306 Ingram, V. M., 280,306 Inokuchi, H., 94, 172 Inoue, H., 60,166 Inouye, M., 283,314 Irwin, D. M., 242,243,281, 282,286, 306 Isaacs, N. W., 24,35, 104, 162, 204, 283, 306,314 Israelachvili,J., 40, 56, 57, 129, 138, 166 Israelachvili,J. N., 56, 166

325

AUTHOR INDEX

Isselbacher, K. J., 25 1, 311 Ivanov, L. V., 74, I71 Iwasa, Y., 60, 165 Iyer, K. S., 270,306 Izumi, T., 60, 166

J Jabbal, I., 250,312 Jack, L., 281,308 Jacoby, W. B., 21,35 Jakobsen, R.J., 109, 166 Jakubowski, H., 147, 168 James, M. N. G., 100, 104, 167,202,307 Janin, J., 117, 172 Jansonius, J. N., 23, 36 Janssen, D. B., 26, 35 JBuregui-Adell, J., 231, 243, 244, 306 Jencks, W. P., 143, 172 Jenkins, J. A, 2, 34 Jenness, R., 189, 197,209, 272,292, 300, 306,311,312 Jensen, L. H., 5,36, 100, 172,216,308 Jernigan, R. L., 119, 120, 162 Jeroszko, J., 208,218,219,260,308 Jigami, Y., 295, 310 Joensson, B., 97, 113, 114, 130, I62 Johansson, C., 288,309 John, C., 29,35 Johnson, J. D., 220,301 Johnson, L. N., 2, 5,20, 26, 33, 174, 175, 193, 195, 198, 199,201, 202,203,216, 229,244,267,283,301,304, 306,311 Johnson, W. C., Jr., 283,294,306 Jollks, J., 175, 181, 182, 229, 231, 240, 241, 242, 243, 244, 245, 246, 282, 283, 284, 285,296,298,305,306,307 Jollks, P., 175, 181, 182, 195, 229, 231, 240,241, 242, 243, 244, 245, 246, 273, 282,283,284, 285,288,296,298, 301, 303,304,305,306,307 Jones, N. D., 31,35 Jones, R., 216,307 Joniau, M., 217, 314 Jordan, F., 297,299 Jorgensen, W. L., 121,167 Jori, G., 215,270,307, 313 Jornvall, H., 189, 227,240, 300 Joynson, M. A,, 194,307

Jukes, T. H., 284,307 Jung, A., 281,307

K Kachalova, G. S., 45, 98, 101, 136, 141, 167,168 Kakalis, L. T., 76, 167 Kakinuma, S., 239, 28 1, 308 Kakudo, M., 283,305 Kalinichenko, L. P., 216, 217, 261, 262, 297,311 Kalk, K. H., 22, 30, 34,35 Kam, Z., 7, 12, 13, 16, 1 9 , 3 4 , 3 5 Kam-Morgan, L. N. W., 275,308 Kaminogawa, S., 227,240,252,307 Kammerling, J. P., 292, 307 Kammerman, S., 181,229,240,240-241, 302 Kanarek, L., 273, 313 Kaneda, M., 231,307 Kang, Y. K., 119,167,294,307 Kaptein, R.,266,301 Karle, I. L., 99, 167 Karlsson, R., 23,36 Karplus, M., 85, 86, 87, 98, 112, 121, 142, 163, 164, 165,167, 168, 169, 170, 171, 194,201,203,205,302,311 Karplus, P. A, 5,34, 106, I 6 7 Kato, I., 231, 307 Kato, S., 259,270,307 Kauffman, D. L., 280,305 Kauzmann, W., 38,40, 41, 42, 43, 45, 50, 54,55,164, 167 Kaverzneva, E. D., 201,309 Kay, C. M.,215,263,264,314 Keegan, R., 197,300 Keenan, T. W., 250,310 Kelders, H. A,, 30, 35 Kell, D. B., 62, 64, 169 Kelly, J. A., 202, 307 Kemp, T. S., 280, 307 Kent, M., 62, 167 Kerby, G. P., 185, 307 Kern, D., 23,35 Kerry, K. R.,292,309 Keshav, S., 241,302 Keszthelllyi, L., 150, 171 Khatra, B. S., 256,257,307

326

AUTHOR INDEX

Khorazo, D., 176, 177,310 Khurgin, Y. I., 43,91,92, 167, 170 Kikuchi, M., 270, 271, 314 Kim, C. Y., 30,35 Kim, K., 50, 167 Kimmel, J. R.,181, 185,313 Kimmich, R.,74, 170 Kimura, M., 277,278,307 King, G., 121, 171 King, L. A., 261, 310 King, N. L. R.,271,301 Kirkpatrick, S., 160, 167 Kirsch,J. F., 201,203,275, 307,308,312 Kirschner, K., 74, 164 Kita, N., 260,307 Kitayama, T., 189,310 Kitchen, B. J., 253, 307 Kiyosawa, I., 189, 255,304, 310 Klee, C. B., 252,254,255,308 Klee, W. A., 252,254,255,270,306,308 Klibanov, A. M., 96, 141, 143, 167, 170 Kliman, P. G., 46,54, 162 Klimanov, A. V., 262,311 Klimova, V. A,, 43, 170 Klotz, I. M., 194, 215, 304, 308 Kluge, A. G., 278,304 Klyachko, N. L., 96, 168 Knight, C. A., 151, 167 Koenig, D. F., 192, 193, 244, 301 Koenig, S. H., 71, 72,76, 167 Koga, K., 213,220,266,308,310 Kollman, P., 121, 167 Kollman, P. A., 121, 162 Kondo, K., 231,245,246,308 Kornberg, A., 296,308 Kossiakoff, A. A., 99, 102, 167, 171, 204, 294,308 Koszelak, S., 29, 34 Krasnopol’skaya, S. A., 76,88, 136, 162 Kravchenko, N. A., 201,309 Krebs,J., 221, 314 Kremer, F., 62,64, 165, 167, 169 Kretsinger, R. H., 216,308 Krigbaum, W. R.,265,308 Kristiansen, K., 229, 241, 314 Kronman, M. J., 187,208,217,218,219, 222, 259,260, 261,262, 263, 264, 267, 297,301,308,312,313 Krummel, B. M., 245,314 Krupyanskii, Y.F., 88,90,91, 130, 136, 137,166,167

Kruse, U., 241,281,311 Kuczera, K., 87, 121, 165, 171 Kugler, F. R.,265,308 Kumagai, I., 239, 281, 308 Kumosinski,T. F., 74, 169,265,311 Kundrot, C. E., 61, 100, 167 Kuntz, I. D., Jr., 38, 40, 41, 42,43,45, 54, 55,56,167 Kurachi, K., 216,308 Kurinov, I. V., 90, 167 Kuwajima, K., 218, 218-219, 220, 260, 265, 269,271,276,297,305,306,307, 308,310 Kydon, D. W., 71,169

L La Rue, J. N., 231,244,245,308 Lafaut, J. P., 262, 314 Laird, J. E., 281,308 Landis, P. L., 31,35 Langer, J. S., 29, 35 Langridge, R.,121,162 Laschtschenko, P., 176, 308 Latovitzki, N., 263, 305 Lavoie, T. B., 275,308 Lawlor, D. P., 183, 197, 310 Leach, S. J., 260,273, 274,278,301,310 Lebedev, Y. O., 269,303 Lebovitz, H. E., 280, 305 Ledieu, M., 229,306 Lee, B., 48,58, 117, 126, 167, 170, 194, 308 Lee, C.-L., 271, 273, 300 Lee, J. C., 60, 167 Lee, J. S., 24,33,35 Lefaucheux, F., 29, 30, 35 Lehmann, M. S., 294,308 Leonis,J., 252,312 Leopold, C., 95, 151, 167 Leung, C. J., 23, 35 Levashov, A. V., 96, 168 Leveque, J. L., 68, 167 Levina, A. A., 76,88, 136, 162 Levitt, M., 87,97, 113, 115, 130, 142, 167 Lewis, M., 189, 209, 210, 211, 212, 213, 239, 247, 248,267, 281, 288, 293, 299, 313 Lewis, P. N., 207,308 Lezina, V. P., 74, 171

AUTHOR INDEX

Liang, P., 22,33 Liao, D.-I., 204, 303 Liberatori, J., 189,302 Lie, 0.. 182, 183, 296, 309 Lienhard, G. E., 215,312 Lieutenant, K., 77, 130, 136, 171 Lifchitz, P., 195, 301 Likhtenshtein, G. I., 74, 76, 88, 136, 162, 167, 171 Lin, M. J., 19, 34 Lin, T.-Y., 271, 309 Lindahl, L., 187,309 Linderstr6m-Lang, K., 294,309 Lindman, B., 72, 76, 166 Ling, G. N., 139, 167 Linse, S., 288, 309 Lioutas, T. S., 75, 168 Lippmann, C., 22,35 Lis, L. J., 58, 168 Littke, W., 26, 29, 34, 35 Litwack, G., 185, 311 Liu, A. K., 231, 244,302 Lonnerdahl, B., 223,309 Lobb, R. R.,298,304 Loftfield, R. B., 147, 168 Longman, R. A., 13, 14, 16, 17,34 Lkonis, J., 208, 264,271,300,309 Looze, Y., 208, 264, 300 Lorber, B., 25,35 Lord, R. C., 262,264,270,271,302,304, 309 Lorder, B., 23,35 Loughnan, M., 189,310 Lovgren, T. N. E., 147, 168 Low, P. S., 147, 168 Luescher, E., 43, 49, 89, 109, 110, 130, 131, 136, 145,164 Luescher, M., 45, 54, 168 Luescher-Mattli, M., 44, 45, 91, 168 Luisi, P., 95, 162 Luisi, P. L., 95, 168 Lumry, R., 112, 147, 168 Lund, E. H., 229,241,314 Lund, W., 42, 172 Luntz, T. L., 295,309

M MacClement, B. A. E., 139, 172 MacGillivray, R. T. A., 227, 238, 309

327

Machin, K. J., 24,35,204,283,306 Machin, P. A., 201,216, 304, 311 MacInnis, J., 122, 168 MacKay, B. J., 182,309 Mackay, D. H. J., 147, 168 Mackay, G. I., 143, 163 Maddox, J., 152,168 Madsen, N. B., 23,34 Maes, E., 208,252,264,300,312 Maes, E. D., 271,309 Magee, S. C., 252,253, 309 Mahk, M.-F., 189, 305 Maher, F., 189, 310 Mainhart, C. R.,274, 275,308,313 Mair, G. A., 174, 192, 193, 198, 203, 212, 213,244,264,266,301 Maksimov, V. I., 201, 309 Malcolm, B. A,, 203, 307 Mallett, C. P., 275, 308 Malmquist, J., 229, 241, 314 Mandelkern, L., 38, 170 Mangelsdorf, I., 241, 281, 303 Mann, G., 178,309 Marden, M. C., 96, 130,162 Margoliash, E., 273, 274, 277, 278, 288-289,295,301,304,309 Margulis, T. N., 100, 172 Mariuzza, R.A., 274,299 Markovic-Housley, Z., 2, 34 Maron, E., 273,300 Maroncelli, M., 122, 168 Marra, J., 40, 56, 57, 129, 138, 166 Martinek, K., 96, 168 Martinez, R.J., 184,313 Masakuni, M., 204,283,306 Mascarenhas, S., 68, 163 Mason, S. A,, 100, 168,294,308 Masui, Y., 8, 34 Matsumura, M., 295,309 Matsuura, Y., 2, 35 Matthew, J. B., 61, 122, 168, 172 Matthews, B. W., 20,33, 104, 105, 166, 172,204,283,295,299,305,309,312, 314 Mattock, P., 180, 305 Maurois, A,, 176, 309 Mawal, R., 191, 255, 304 McAlister, M., 58, 168 McCammon, J. A., 87,98,163, 168 McCanny, J., 68, 171 McDonald, B. L., 189,310

328

AUTHOR INDEX

McDonald, C. C., 215,265,309 McGuire, E. J., 150, 312 McGuire, W. L., 191,309 McIntyre, G. J., 100, 168, 294,308 McKeever, B. M., 29,34 McKenzie, H. A., 40,41,99, 117, 118, 164, 178, 182, 183, 184, 187, 189, 190, 191, 194,204,214,222, 223,224,227, 229, 232,238,240,241, 243,247,260, 263, 264,268,272,287,291,292,293, 296,299,300,301,303,304,305,306, 307,309,314 McLachlan, A. D., 118, 142, 164 McLaren, A. D., 41,42,44,94, 168, 171 McLaughlin, P. J., 283,306 McLean, V. E. R.,62, 166 McMurry, S., 181, 302 McPhee, M. S., 251,311 McPherson, A., 2, 7,21, 26, 29,33-34, 34,35 McPherson, A. A,, 30.35 Medvedeva, N. V., 91,92, 167 Meehan, E. J., 22, 26, 29,33-34,35 Meehan, E. J., Jr., 21, 34 Mejdoub, H., 23, 35 Mendelsohn, R.,264,302 MenCndez-Arias, L., 231, 246, 273, 288, 304,312 Mercier, J.-C., 227, 232,239, 281, 304, 314 Mercier, J. C., 189, 305 Merlin, L. M., 250,310 Messer, M., 189,292,307, 309, 310 Metschnikoff, E., 176, 310 Meyer, E., 105, 144,168 Meyer, K., 176, 177,310 Meyer, W., 62, 167 Mezei, M., 120, 121, 168 Michael, J. G., 273, 274, 278, 301 Michel, H., 2, 35 Middlendorf, H. D., 85,86, 133, 168 Migliardo, P., 110, 162 Mikol, V., 21, 23, 25, 32, 34, 35 Mille, M., 150, 168 Miller, A., 273, 274, 278,301 Miller, J. N., 261, 310 Mills, S. E., 21, 34 Milos, M., 218, 312 Mitani, M., 297,310 Mitchell, G., 273, 314

Mitranic, M. M., 191, 252, 310 Mitschler, A., 25, 35 Miura, K., 239,281,308 Mizobuchi, H., 252, 307 Moessbauer, R. L., 90, 130, 167 Mol, C. D., 23,24,33 Mollenhauer, H. H., 250, 303 Molnar, R.E., 278,300 Mornany, F. A., 208,210,266,271,314 Monizingo, A. F., 105, 166 Montagu, M. V., 152, 172 Mookerjea, S., 252, 304 Moor, U., 52,54, 170 Moras, D., 23,25,32,34,35,41, 168 Mordick, T., 255,257,310 Morgan, F. J., 181, 229, 231, 240, 240-241,300,302,313 Morgan, H., 66,68, 162, 168 Mori, Y., 94, 172 Morikawa, M., 295, 310 Morita, Y., 26, 35 Morowitz, H. J., 42, 172 Morozov, V. N., 43,45, 98, 101, 136, 141, 165,167, 168 Morozova, L.A., 216,217,221,261,297, 311 Morozova, T. Y., 45,98, 101, 136, 141, 167,168 Morrt, D. J., 250,310 Morris, D. W., 30, 35 Morrison, J. F., 256, 257, 310 Morsky, P., 186, 310 Moscarello, M. A,, 191, 252, 310 Moser, I., 185, 310 Mossop, G. S., 292, 310 Moult, J., 100, 115, 116, 142, 166, 194, 197,310 Mouton, A., 231,241, 245,307 Mrevlishvili, G. M., 50, 54, 162, 168 Mross, G. A., 229, 231,241,244,287,288, 311,314 Muchrnore, S. W., 8,33 Mukhin, E. N., 76, 88, 136, 162 Muller, V., 189, 292,301 Muller, V. L., 227,229, 292, 309 Mullinax, F., 191,306 Multon, J. L., 152, 171 Municio, A. M., 283, 304 Munjal, D. D., 189, 251, 312 Munks, S., 189, 310

329

AUTHOR INDEX

Murakami, K., 217,219,262,266,297, 301, 310, 311 Muraki, M., 295,310 Murphy, W. H., 187, 189,227,232,238, 300,301 Murthy, K., 24,35 Musci, G., 220, 223, 266,310 Mussig, J., 21,'24, 36 Myachin, E. T., 101, I 6 7 Myachin, T. T., 45, 98, 136, 141, 168

212,213,229,244, 255, 264, 266, 271, 301,302,306,307 Norton, R. S., 266, 301 Now, B. T., 23,35 Nowik, I., 88, 162 Nusser, W., 74, 170 Nylander, T., 59, 164 Nyquist, S. E., 250, 303

0

N Nagabhushan, T. L., 29,34 Nagamatsu, Y., 189,310 Nagasawa, T., 189, 310 Nagel, R. L., 19, 34 Nagle, J. F., 150, I68 Nagy, J. A,, 142,172 Nakanishi, M., 265,271,313 Nakao, M., 270,271,314 Napolitano, L., 189, 229, 241, 302, 304 Narita, K., 231, 307 Nascimento, 0. R., 77, 136, 170 Naumann, R., 13, 14, 16,26,29,3?-34, 35 Navia, M. A., 29, ?4 Nelson, B., 26, 29,33-34 Nelson, G., 29, 34 Nemethy,G., 117, 118, 119, 142, 167, 168, I69 Nkmethy, G., 294,307 Nicholas, K., 189, 310 Nicholas, K. R., 189,310 Nicola, N. A., 260, 310 Nicolle, M., 176, 310 Nielsen, S. O., 80, 166 Nienhaus, G. U., 88,89, 136,169 Nightingale, N. R. V., 62, 166 Nihoul-Deconinck, C., 273,313 Nilsson, L., 41, 164 Nitta, K., 218, 222, 223, 260, 269, 288, 289,290,297,303,305,307,308,310, 313

Noble, M., 273, 314 Noda, Y., 52,60, 140, 165 Norman, J. A,, 97,98, 171 North, A. C. T., 174, 175, 192, 193, 194, 198,201, 202, 203,206,207, 208, 210,

Oatley, S. J., 294, 300 Ochman, H., 278,314 Oefner, C., 105, 144, 169 Offord, R. E., 231,244,312 Ohlendorf, D. H., 26,29,34,36 Oka, T., 189, 250, 310 Okamura, M., 270,307 OKeeffe, E. T., 253,254,255,257,310 Ollis, D., 2, 25, 33, 35 Ollis, D. L., 24, 35 Olsen, K. W., 251,300 Ondrias, M. R., 110, 165 Ono, M., 250,310 Ono, T., 216, 306 Oobatake, M., 118, 119, 142, 169 Ooi, T., 118, 119, 142, 169 Orbell, J. D., 24, 35 Ord, V. A., 74, 75, 165 Osserman, E. F., 175, 182, 183, 197,229, 241,287,288, 309,310, ?I4 Ostrovsky, A. V., 262, 311 Otsuka, A. J., 20, 33 Otting, G., 73, 169 Ovsepyan, A. M., 97, 111, 131, 140, 148, 163, 164 Owen, J., 283,312

P Packer, L., 41, I69 Padlan, E. A,, 105, 144, 171,274,313 Page, M. I., 143,162 Pahler, A., 8, 35 Pal, G. P., 105, 162 Pal, P. K., 111, I71 Pallansch, M. J., 46, 54, 162 Palmer, J. W., 176, 177, 310

330

AUTHOR INDEX

Palmer, K. J., 197,311 Palmer, R. A., 20,35 Pantin, V. I., 96, 168 Parak, F., 62, 64, 88, 89, 90, 91, 103, 130, 136, 148,165,166,167,169,171 Parello, J., 74, 163 Parker, D., 281,299,302, 303,305,308 Parrym, R. M., 183,302 Parsegian, V. A., 40, 56, 57, 58, 129, 138, 150,168, 169, 170,172 Paslay, J. W., 8, 33 Pastuszyn, A., 147, 168 Patrono, D., 182, 302 Pauling, L., 277, 284, 315 Pauly, H., 50, 162 Pearce, R. J., 183, 187,296, 314 Pedersen, K. O., 178,311 Peemoeller, H., 71, 169 Peer, W. J.. 117, 168 Perkins, H. R., 177,311 Perkins, S. J., 199, 216, 267,311 Permyakov, E. A., 85, 169, 216, 217,221, 261,262,297,311 Perry, A. L., 2 0 , 3 5 Perutz, M. F., 61, 169 Pessen, H., 74, 169, 265, 311 Peters, C . W. B., 241, 281, 311 Peters, D., 103, 169 Peters, J., 103, 169 Pethig, R., 42,43, 62, 64, 66, 68, 162, 163, 164,165,166,168, I69 Petrozzo, M. A,, 31, 36 Petry, W., 87, 164 Petsko, G. A,, 198, 303 Pfeil, W., 272, 311 Phillips, D. C., 174, 175, 189, 192, 193, 197, 198, 199, 201, 202, 203, 209, 209, 210, 211, 212, 213, 216, 223, 229, 231, 244, 247, 248, 264,266, 267, 281, 283, 288, 293, 294,299,300,301, 304, 306, 307,31I , 313 Phillips, G. N., Jr., 22, 33 Phillips, N. I., 189, 272, 311, 312 Phillips, S. E. V., 103, 169, 274, 299 Phillips, W. D., 215, 265, 309 Piculell, L., 72, 169 Pierce, M., 250, 311 Pincus, M. R., 202,311, 313 Pintar, M. M., 71, 169 Pinteric, L., 250, 312 Pissis, P., 68, 69, 162, 167

Pittner, F., 185, 310 Podjarny, A., 194, 197,310 Podolsky, D. K., 250,251,311 Poglitsch, A., 62,64, 165, 167, 169 Pogolotti, A., 16, 17, 33 Poljak, R. J., 174, 192,274,299,301 Pollock,J. J., 182, 309 Pollwein, R., 241, 281, 311 Polnaszek, C. F., 73, 97, 128, 169 Ponnuswamy, P. K., 118, I70 Poole, P., 87, 171 Poole, P. L., 42, 55, 83, 108, 109, 122, 162, 165,169 Post, C . B., 112, 169, 194,201,203,311 Pottel, H., 271, 305 Potter, M., 274,275, 313 Potthast, K., 94, 169 Poulsen, F. M., 80, 164, 213,266, 311 Powell, J. T., 250, 252, 253, 254, 256, 311 Powell, K., 29, 34 Powning, R. F., 181, 311 Prabhakaran, M., 118, 170 Prager, E. M., 181, 185, 229,231, 242, 243, 244, 245, 246,273, 274, 275, 278, 284,285,286-287,289,290,296,301, 303,306,307,308,311,313,314 Prasad, L., 24,35 Prasad, R., 185, 240,311 Prasad, R. V., 189, 227, 240,311 Prieels, J.-P., 190, 208, 252, 257, 264, 272, 300,312 Prieur, D. J., 284, 312 Privalov, P. L., 54, 168 Proctor, C. M., 185,303 Proctor, S. D., 189, 190,312 Proctor, V. A,, 175, 312 Prouty, M. S., 58, 129, 170 Ptitsyn, 0. B., 269,276,294,303,304,312 Puchwein, G., 74, 164 Pulford, W. C. A., 99, 100, 101, 102, 104, 127, 144, 146,163 Pulsinelli, P. D., 110, 163 Pusey, M., 26, 29, 33-34 Pusey, M. L., 13, 14, 16,35

Q Qasba, P. K., 189,240, 246, 281,303,312 Quarfoth, G. J., 189,312 Quiocho, F. A., 105, 170

33 1

AUTHOR INDEX

R Rabinovich, D., 194, 197, 310 Radhakrishnan, R., 105, 144,168 Radzicka, A., 118, 170 Rahman, A., 112,165 Ralston, G. B., 292, 310 Ram, B. P., 189,251,312 Rand, R. P., 40, 56, 57, 58, 129, 138, 150, 168,169,170 Randall, J. T., 85,86, 168 Ranghino, G., 120, 164 Rao, K. R., 223,269,288,312,315 Rao, K. S., 44, 170 Rao, P. B.,42, 81, 84, 162, 170 Rao, S., 121, 167 Rau, D. C., 40, 56, 57,58, 129, 138, 169, 170 Ravichandran, V., 99, 171 Rawitch, A. B., 261, 265,312 Raymond, J. A,, 151, 170 Reardon, I. M., 8, 33 Rees, A. R., 231, 244,312 Rees, G. D., 95-96,165 Reichlin, M., 273, 274, 278, 301 Reinbolt, J., 23, 35 Reinisch, L., 88, 90, 96, 130, 162, 169 Remington, S. J., 204, 283,303,309,312, 314 Renkawitz, R., 241, 281, 303 Rennekamp, G., 103, 169 Ressler, N., 110, 147, I71 Reuscher, H., 103, 166, 169 Reyerson, L. H., 45, 112, 127, 165, 166, 168 Reynolds, A. H., 96, 130, 162 Reynolds, C. D., 104, 162 Rice, D. W., 194,300 Richards, F. M., 00, 48, 50, 5 1, 6 1, 117, 126, 141,167,170, 194,308 Richmond, T. J., 117, 170 Ries-Kautt, M. M., 197, 312 Rigler, R., 41, 164 Ritchie, A., 278, 300 Rizvi, T. Z., 71, 162 Robbins, F. M., 208, 260,261, 264,267, 308,312 Robert, M. C., 29, 30, 35 Robinson, R., 176,209 Robinson, R. D., 257,301 Robinson, R. H., 95-96, 165

Rochester, C. H., 42, 170 Rodeau, J.-L., 2 1 , 3 5 Rodeau, J. L., 23,32,34 Rodriguez, R., 231, 246, 273, 283, 288, 304,312 Rogers, C., 189,301 Rose, J. P., 22, 35 Roseman, S., 250,312 Rosenberg, A., 80,97,98,166, 170,171 Rosenberg, S., 201,312 Rosenberger, F., 2, 16, 21, 22,32,34,35 Roslyakov, V. Y., 91, 92, 167, 170 Ross, P. D., 8, 12, 18, 34, 35 Rossky, P. J., 112, 167, 170 Roth, S., 250, 311 Rottenberg, M., 45, 54, 168 Rowe, T., 278, 304 Rowen, J. W., 41,42, 168 Rozeboom, H. J., 26,35 Ruani, G., 68, 163 Ruegg, M., 44,45, 52, 54, 91, 168, 170 Ruff, M., 20, 25, 3 4 , 3 5 Ruggiero, J., 77, 136, 170 Rumball, S. V., 208, 210,266, 271, 314 Rupley, J. A., 16, 17,33, 39, 44, 45, 47, 48, 49, 51, 64, 65,66,67, 69, 77, 78, 81, 82, 83, 92, 93, 100, 108, 111, 112, 122, 125, 126, 127, 131, 133, 135, 140, 141, 142, 144, 145, 150,162, 163,170, 171,172, 175, 193, 198, 201, 202, 229, 244, 260, 263,300,302,306,312,313 Russell, A. J., 96, 170 Russell, S. T., 122, 171 Rydstedt, L., 273,300

S Sable, H. Z., 215, 266, 304 Saccomani, G., 270,271, 302,313 Saenger, W.,41,99, 105, 162, I70 Safaya, S. K., 246, 281, 312 Sage,G.W., 208,218,219,260,308 Sakabe, K., 104,162 Sakabe, N., 104, 162 Sakar, P. K., 272,312 Salemme, F. R., 5, 16, 17, 22, 26, 29,34, 35,36, 104,165 Salton, M. R. J., 177, 178, 312 Salunke, D. M., 5, 33, 104, 163 Samanta, S. R., 110, 170

332

AUTHOR INDEX

Sanches, R.,77, 136, 170 Sander, C., 87, 167 Sarma, R., 202,283,305,312 Sarrna, V., 194,307 Sarrna, V. R.,174, 192, 193, 198,203,212, 213,244,264,266, 301 Saroff, D. A,, 275,308 Sasabe, H., 2,34 Savage, H., 99, 106,170 Sawyer, W. H., 187,300 Sax, M., 22,35, 151, 172 Saya, A., 194, 197,310 Scanlon, W. J., 61, I70 Scatturin, A., 270,313 Schachter, H., 250, 312 Schaer,J.-J, 218, 312 Schanbacher, F. L., 256,312 Schauer, G., 74,170 Schauer, R.,292,307 Schechter, A. N., 58, 129,170 Schejter, A., 110, 162,295,309 Scher, H., 156, 157,170,172 Scheraga, H. A., 38, 117,118, 119, 142, 167, 168, 169, 170, 202,207,208, 210, 266,269,271,294,300,304,307,308, 311,312,313,314 Schilling,J. W., 275, 284, 308, 313 Schillinger,W. E., 72, 167 Schindler, M., 202,257,272,312 Schindler, P., 45,54, 168 Schinkel,J. E., 51, 81, 82, 83, 140, 141, 170 Schlichtkrull,J., 13, 14, 16,36 Schlitter,J., 77, 130, 136, 171 Schlusselberg,J., 190, 312 Schmidt, D. V., 189,227,312 Schoenborn, B. P., 99, 103,166,170 Schoentgen, F., 181,229,231,242, 245, 246,282,284,307 Schreiner, L. J., 71, 169 Schrier, E. E., 46, 126, 127, 162 Schuler, L. A,, 232, 281,306 Schulz, G. E., 5, 34, 106, 167 Schuster, I., 74, 164 Schutz, G., 281,307 Scoffone, E., 215,307 Scordamaglia,R., 120, 164 Sebelien,J., 178, 312 Secemski, I. I., 215,312 Segawa, T., 218,297,313

Seheshadri, B. S., 2,36 Seibel, G., 121, I67 Sekharudu, Y. C., 99,171 Selsted, M. E., 184,313 Sernisotnov,G. V., 269,276,294,303,304 Sen, A., 197,272,300,312 Senadhi, V., 29, 34 Sendhi, S. E., 29,34 Sercarz, E. E., 273, 274,278,301 Shablakh, M., 68, 170 Shahani, K. M., 183,302 Shakhnovich, E. I., 269, 312, 313 Shaper, J. H., 251,300 Sharma, R. N., 271,313 Sharon, N., 202,215,257,263, 264,272, 312,314 Sharon, R., 97, 113, 115, 130, 142, 167 Sharp, A. R., 71, 169 Shaw, D., 189,310 Shaw, D. C., 182, 183, 187, 189,222,227, 229,232, 238,240, 241,243, 247,287, 291,292,296,301,305,307,309, 310, 314 Shchegoleva,T. Y., 64, 170 Sheats, G. F., 85, 170 Shenoy, R. K., 71,169 Sheriff, S., 5, 36, 274, 275, 308, 313 Sherman, F. B., 43, 170 Shewale,J. G., 189, 227, 232, 238, 240, 248,287,313 Shimamoto, N., 259,307 Shirnanovskii,N. L., 74, 171 Shimazaki, K., 222, 288, 297,310 Shirley, W. M., 71, 163, 171 Shlichta, P., 7, 35 Shoham, M., 106, I71 Shore, H. B., 7, 12, 13, 16, 19, 35 Shporer, M., 72, 167 Shrake, A., 48, 126, 171, 198,260,306, 313

Shugar, D., 185,313 Sidow, A., 281, 306 Sieker, L. C., 5, 36, 100, 172, 216,308 Sielecki, A. R., 100, 104, 167, 202, 307 Siernankowski, L., 66,67,69, 142, 144, 150,170 Silverton, E. W., 274,313 Silvia,J. C., 253, 302 Silvia,J. D., 253, 313 Simatos, D., 152, 171

AUTHOR INDEX

Simmons, E. R.,253, 313 Simmons, N. S., 263, 304 Simons, P., 110, 165,298, 314 Simpson, R.J., 181,231,313 Singh, G. P., 62,64,89, 130, 136, 171 Singh, U. C., 121, 162, 167 Sinha, S. K., 189, 217, 218,219,227, 232, 238,240,248,262,287,297,308,313 Sippel, A. E., 241, 281, 307, 311 Sjoelin, L., 104, 172 Sjogren, B., 178,313 Skujins, J. J., 94, 171 Sledziewski, A,, 241, 302 Sloan, D. L., 74, 171 Smilansky, A,, 194, 197, 310 Smille, L. B., 280, 305 Smith, C. D., 29, 34 Smith, C. W., 105, 144, 171 Smith, E. L., 181, 185, 313 Smith, G. N., 182, 313 Smith, H. W., 29, 34 Smith, J., 85, 86, 87, 164, 171 Smith, J. L., 24, 33 Smith, K. B., 109, 166 Smith, R.,20, 34 Smith, S. G., 189, 197,209, 210, 213, 248, 288,300,313 Smith-Gill, S. J., 202, 273,274,275, 278, 301,308,313 Smolelis, A. N., 185, 313 Snaith,J. W., 50, 54, 166 Snyder, R.,26, 29,33-34 Snyder, R.S., 13, 14, 16,33,35 Sobel, J., 273, 300 Sobel, J. H., 181, 183, 229, 240, 240-241, 302 Sokhadze, V. M., 50, 162 Solbakken, A,, 112,168 Solbu, H., 183, 309 Somero, G. N., 147, 168 Sommers, P. B., 261, 313 Somogyi, B., 97,98, 170,171 Sophianopoulos, A. J., 267, 306, 313 Sorensen, L. B., 96, 130, 162 Serensen, M., 178,313 Serensen, S. P. L., 178, 313 Soulier, S., 227, 232,239, 281,304, 314 Southgate, C. C. B., 118, 172 Southworth, M. W., 152, I72 Spangler, B. D., 24, 36

333

Speck, J. C., 231,244,245,308 Spencer, R., 2 1 , 3 5 Spevak, W., 241,302 Sproule, R. C., 68, 171 Stauffer, D., 66, 156, 157, 171 Steigemann, W., 103, 166, 169 Steinberg, M. P., 75, 168 Steinhoff, H. J., 77, 130, 136, 171 Steinman, H. M., 180, 189, 292,302, 305 Steinmann-Hofmann, B., 95, 168 Steinrauf, L. K., 194, 197,307,313 Steitz, T. A., 21, 24, 25, 33, 35, 36 Stepanyants, A. U., 74, 171 Stern, P. S., 87, 167 Sternberg, M. J. E., 294,300 Stevens, E., 94, 171 Stevens, L., 94, I71 Stewart, C.-B., 229,242, 243,284,285, 307,313 Stillinger, F. H., 112, 165 Stocker, C., 182,313 Stockmayer, W. H., 154, 171 Stonehouse, J. R.,60, 171 Stout, J. W., 49, 166 Strambini, G. B., 84, 96, 166, 171 Strandberg, B., 41, 168 Strokopytov, B. V., 101, 167 Strosberg, A. D., 273,313 Strydom, D. J., 298,304 Stuart, D. I., 189, 193, 209, 210, 211, 212, 213, 223,239,247, 248, 267,281,288, 293,299,311,313 Stura, E., 23,36 Sturtevant, J. M., 60, 171 Suck, D.,41, 105,144,168,169 Suddath, F. L., 26,29,33-34 Sugai, S., 218, 220, 221, 222, 223, 260, 265, 269, 271,288,289,290, 297,303, 305,306,307,308,310,313 Suguna, K., 105, 144, 171 Sullivan, J., 112, 168 Sundaralingam, M., 99, 171, 209,210,313 Sussman, F., 121, 172 Sussman, J. L., 106, 171 Sutcliffe, J. W., 110, 163 Suurkuusk, J., 49, 126, 171 Suzdalev, I. P., 90, 130, 167 Svedberg, T., 178,313 Svensson, L. A., 104, 172 Swan, I. D. A., 197,265,299,300

334

AUTHOR INDEX

Swarte, M. B., 24, 36 Swartzenderber, J. K., 3 1 , 3 5 Swift, T. J., 215, 266, 304 Syed, M., 182, 183,296,309 Sykes, B. D., 202,307 Synder, R. E., 29,34 Szent-gyorgyi, A., 66, 165

T Tabak, M., 77, 136,170 Tabony, J., 95-96,165 Takano, T., 103, I71 Takesada, H., 265, 271, 313 Takizawa, T., 17, 36 Tamaki, E., 239,281,308 Tamburro, A. M., 270,271,302,313 Tammers, D., 29,34 Tanahashi, N., 179,272,302,313 Tanaka, F., 111, 171 Tanaka, H., 295,310 Tanaka, S., 8, 18, 26, 33, 68, 110, 162 Tanford, C., 270,313 Taniyama, Y., 270,271,314 Tapia, O., 147, 171 Taylor, C. A., 20, 34 Taylor, G., 29, 34 Teahan, C. G., 182, 189,222,223,229, 243,246,247,287,291,299,305,314 Teeter, M. M., 100, 102, 166, 171 Teichberg, V. I., 215,263, 264, 314 Teleman, O., 97, 113, 114, 130, 162 Teller, E., 43, 163 Ten Eyck, L. F., 283,312 Tessier, J. H., 189, 305 Thaller, C., 23, 36 Thanki, N., 99, 171 Theriault, N. Y., 8, 33 Thierry, J. C., 23, 25, 32,34, 35 Thompson, E. 0. P., 181, 185,292,313, 314 Thompson, K., 273,314 Thompson, M. P., 197,209,300 Thompson, R., 176, 177,310 Thomsen, J., 229,241,314 Thornton, J. M., 99, 162, 171 Thulin, E., 288,309 Tidor,B., 87, 121, 164, 165, 171 Tiktopulo, E. I., 269, 303

Tiller, W. A,, 19, 32, 36 Timasheff, S. N., 2 , 5 , 6 , 3 3 , 3 6 , 4 1 , 6 0 , 166,167,265,311 Ting, K. L., 119, 120, 162 Titani, K., 231, 307 Tjian, R., 201, 304 Todd, P. E., 273,274,278,301 Tollin, G., 77,78, 92, 93, 133, 170 Tombs, M. P., 189,301 Tomich, C.-S. C., 8, 33 Tominaga, N., 231,307 Tracey, D. E., 8 , 3 3 Traub, W., 194, 197,310 Trayer, I. P., 180,251,305,314 Treacy, G. B., 227,229, 291,292,306, 309 Tredgold, R. H., 68, 171 Treffry, A., 88, 162 Trewhella, J., 85, 86, 164 Tronrud, D. E., 105, 166 Tsuboi, M., 265,271,313 Tsuda, M., 94, 172 Tsuge, H., 222,223,288,289,290,297, 310,313 Tsugita, A., 283, 314 Tulinsky, A., 105, 146, 163 Turkington, R. W., 250, 314 Turley, E. A,, 250,311 Turner, A,, 143,163 Tusupkaliev, U., 43, 170 Tuttle, R. W., 208, 210, 266, 271, 314 Twigg, P. J., 2 1.34

U Uechi, M., 189, 310 Ullrich, V., 80, 164 Usha, M. G., 74, 136, 171 Utiyama, H., 259, 270, 307

v Vacatello, M., 115, 117, 142, 166 Vadali, G., 270, 313 Vagin, A. A., 101, 167 Valitov, V. M., 62, 165 Vallee, B. L., 298,304 Van Cauwelaert, F., 217, 220, 221, 223, 261,262,271,303,305,314

AUTHOR INDEX

Van Ceunebroeck, J.-C., 217,221,314 Van Dael, H., 223,262,303,314 Van Der Laan, J. M., 24,36 van Halbeek, H., 292,307 Van Holde, K. E., 267, 313 Van Leemputten, E., 231,241, 245,307 Vanaman, T. C., 179, 180, 189,206,207, 208,210,227,232,250,255,266, 271, 302,305,314 Vandekerckhove, J., 152,172 Vander Meulen, D. L., 110, 147, 171 Vandermaelen, P. J., 22,34 Vandonselaar, M., 24, 35 Varo, G., 150, 171 Vasu, S., 20,33 Vecli, A., 68, 150, 163 Velicelebi, G., 60, 171 Velick, S. F., 74, 171 Venkatappa, M. P., 2 , 3 6 Venyaminov, S. Y., 276,294,303 Vernon, C. A., 200,314 Vijayan, N. M., 104, 162 Vilotte, J.-L., 227, 232,239, 281, 304, 314 Vincentelli, J. B., 271, 309 Vitols, R., 267, 308 Vliegenthart, J. F. G., 291,307 Vogel, H. J., 187,309 Vol'kenshtein, M. V., 101, 167 von Bahr-Lindstrom, H., 189,227,240, 300 Vonderhaar, B. K., 189,301 Vyas, N. K., 105, 170

W Wadden P., 252,304 Walker, N. P. C., 189,209,210,211, 212, 213,239,247, 248,267, 281,288, 293, 299,313 Walrafen, G. E., 110, 170 Walton, A. R., 55, 169 Wanderlingh, F., 110, 162 Wang, B.-C., 22,35 Ward, K. B., 3 1 , 3 6 Warme,P. K., 208,210,266,271,314 Warner, R. C., 182,303 Warren, G. J., 152, 172 Warshaw, A. L., 251,311 Warshel, A., 121, 122, 143, 164, 171, 172

335

Wasacz, F. M., 109, 166 Watenpaugh, K. D., 8 , 3 3 , 100, 172 Watkins, W. M., 178,314 Watt, I. C., 41-42, 172 Weaver, D. L., 142, 167 Weaver, L. H., 23,36, 104, 105, 166, 172, 204,283,305,314 Weber, P. C., 8,22,26, 27, 29,30,33,34, 36, 104, 165 Wedel, A., 241, 281,303 Weiser, M. M., 250,311, 314 Weiser, R. S., 178, 301 Weiss, R. M., 121, 171 Weissman, L. S., 245,314 Welberry, T. R., 20,34 Welch, G. R., 41, 172 Wendoloski, J. J., 122, 172 Werber, M. M., 96, 130, 165 Westbrook, E. M., 24,36 Westerman, A. V., 42, 170 Wheelcock, J. V., 189, 190, 312 Whitaker, J. R., 182, 314 White, F. H., Jr., 183, 184, 187, 214, 222, 270,271,272,293,296,309,314 White, R. T., 28 1, 306 White, S., 2, 35 White, T. J., 229, 231, 241, 244, 277, 286, 287,288,306,314 Whitlow, M. D., 102, 171 Whittaker, R. G., 292,314 Wichmann, A., 178,314 Wiggens, P. M., 139, 172 Wilcox, P. E., 180, 315 Wilkinson, A., 42, 172 Williams, J. W., 267, 303 Williams, R. J. P., 216, 303 Wills, P. R., 60, 172 Wilmut, I., 298, 314 Wilson, A. C., 181, 183, 185, 203, 229, 231, 241, 242,243, 244,245, 246, 273, 274, 275, 277,278, 281,282, 284, 285, 286,286-287,287,288,289,290,296, 301,303,304,305,306,307,308,311, 313,314 Wilson, D. K., 105, 170 Wilson, E., 23,36 Wilson, I. A., 209, 210, 313 Wilson, J. R., 250, 314 Wilson, K., 22,35,41, 168,295,299 Wilson, K. R., 147, 168

336

AUTHOR INDEX

Wilson, K. S., 194,300 Winborne, E. L., 8 , 3 4 Winzor, D. J., 60, 172 Witherow, W., 13, 16,35 Witherow, W. K., 16,33 Wittebort, R. J., 74, 136, I71 Wittmann, H. G., 21, 24,36 Wlodawer, A., 99, 104, 106, 170, 172 Wodak, S. J., 117, 172 Wolber, P. K., 152, 172 Wolfenden, R., 18, 26,33, 118, 170, 172 Wolynes, P. G., 98, 168 Wong, C. H., 143,172 Wong, L.-J. C., 253, 314 Wong, S. S., 253, 314 Woods, K. L., 250,315 Woods, K. R., 189,227,239,273,306 Woodward, C. K., SO, 172 Wozniak, J. A,, 295, 299 Wright, A. G., Jr., 270,271, 314 Wright, C. S., 105, I72 Wuethrich, K., 73, 169 Wyckoff, H. W., 2, 36

X Xanthopoulos, K. G., 282-283,303 Xuong, J., 2 1 , 3 4

Y Yagi, T., 94, 172 Yamada, H., 216,306 Yamaguchi, K., 252,307 Yaniamoto, Y., 270, 271, 314 Yang, D., 22,35 Yang, D. S. C., 151, 172 Yang, J. T., 264,302 Yang, P., 45,47, 48,49, 100, 126, 127, 131, 172

Yang, P.-H., 263, 302 Yang, P. H., 66, 77, 78,92, 93, 108, 112, 122, 133, 142, 144, 150,163,170 Yaparidze, G. S., 50, 162 Yarmolenko, V. V., 216,217, 261,297,311 Yasonubu, K. T., 180,315 Yathindra, N., 99, 171 Yeh, Y., 151, 165 Yem, A. W., 8,33 Yeomans, F. G., 71, 169 Yonath, A., 21,24,36, 194, 197,310 Yoneyama, M., 269,308 Yoo, C. s.,22,35 Yoshimura, Y., 60, 166 Yost, V., 26, 29, 33-34 Young, C. C., 19, 32,36 Young, P. R., 143, 172 Young, R. D., 148, 165 Yu, N.-T., 262, 309, 315 Yue, K. T., 96, 130, 162

z Zaidi, Z. H., 189, 227, 240, ?OO Zaks, A., 96, 141, 143, 172 Zallen, R., 66, 67, 156, 157, 158, 159, 164, 170, 172 Zambrowicz, 22,33 Zempel, L., 97.98, 171 Zeng, J., 288, 315 Zhang, J., 55, 169 Zhang, L., 203,307 Zientara, G. P., 142, I72 Zimmerberg, J., 150, 172 Zuckerkandl, E., 277,284,315 Zuckerman, J. M.,182, 309 Zuk, W. M., 31,36 Zurcher-Neely, H. A., 8, ?3

SUBJECT INDEX A Ab znilio simulations, hydration, 120 Absorbance spectroscopy, hydration, 110-111 Activation energy, heterogeneous nucleation, 7 Active-site waters, 104- 106 Aggregation, a-lactalbumin and lysozyme, 267-268 Albumin critical hydration level, 76-77 crystals, hydration dependence of volume, 50-51 fluorescence, 85 water proton relaxation, 73 Amide hydrailon and, 107-109 hydrogen exchange, 80-82,97-98, 135 Amino acid, composition a-lactalbumin, 224-227, 232 lysozyme, 224-225, 228-232 Anhydrobiosis, 151 Antifreeze proteins, 151- 152 Antigenic determinants a-lactalbumin, 273-274 lysozyme, 273-274 L-Arabinose-binding protein, water role in binding, 105 Association, a-lactalbumin and lysozyme, 267-268 Association constants, Ca(I1) binding to a-lactalbumin and lysozyme, 297-298

B Bacteriolysis, a-lactalbumin, 176- 177 p proteins, water binding to carbonyl groups, 109

Bound water, see Hydration shell Brunauer-Emmett-Teller theory, 43

C Ca(I1) a-lactalbumin binding, 2 16-2 18, 220-221,248-249,288 association constants, 297-298 binding site, 2 13 lysozyme binding, 216, 222,248-249 association constants, 297-298 Capacitance, hydration dependence, 64 -66 Carboxylate IR bands, hydration and, 107-109 Casein, depolarization current band, 68-69 Catalysis, fluctuations and, 148- 149 Cell lysis, kinetics, lysozyme, 183- 185 Chymotrypsin active site, 146 acylation, 91-92 enzyme activity, 91-92 a-Chymotrypsin, critical hydration level, 76-77 Chymotrypsinogen A, tryptophan lifetime, 85-86 Circular dichroism hydration, 11 1 a-lactalbumin and lysozyme, 263-265 Compressibility, hydration, 6 1 Conformation, hydration, 139- 141 Connectivity, 153 Cow milk, lysozyme isolation, 182- 183 Crarnbin, diffraction, 102- 103 Crystallization, 1-32 automated, 30-31 cessation of growth and crystal disorder, 19-20

337

338

SUBJECT INDEX

Crystallization (continued) competition between nucleation and growth, 17-19 conditions, searching for, 25-28 density gradients, 29 driving forces, 3-6 growth mechanisms controlled by surface kinetics, 13- 14 growth rate measurement, 14- 15 molecular preassociation role, 16- 17 transport-controlled, 13 transport phenomena, 15- 16 growth unit, 4-5 hanging-drop technique, 30 heterogeneous nucleation, 7- 12 homogeneous nucleation, 6- 7 methods achieving different conditions for nucleation and growth, 22 batch, 20 dialysis, 20-21 free interface diffusion, 22 seeding, 23 temperature shift, 2 1-22 vapor diffusion, 2 1 in microgravity, 26,29-30 nucleation rate, 7-8, 12 precipitants, 5 protein purity, 23-25 solution turbulence, 29 stages, 3 using successive automated grid searches, 26-28 Crystals disorder, cessation, 19-20 destabilization, 5 poisoning of, 19 structural defects, 19-20 Cyclohexane, transfer free energies, 118 Cysteine, oxidation, 24

D Denaturation hydration, 52-53 a-lactalbumin and lysozyme, 268-27 1 Density gradients, crystallization, 29 Dialysis, crystallization, 20-2 1 Dielectric relaxation, hydration, 61-69

Diffraction hydration, 99- 107 nonfreezing water, 55 Disulfide bridges, a-lactalbumin and lysozyme, 247 Domain coalescence, protein folding, 142

E Electron spin resonance hydration, 76-79 a-lactalbumin and lysozyme, 265-267 Electrostatic simulations, hydration, 122 Empirical valence bond method, 121 Enthalpy, hydration, 45-46 Enzyme activity chymotrypsin, 91-92 hydration, 135, 144 Excluded volume model, 60 Exons, a-lactalbumin and lysozyme, 280-282

F Fab fragments, 24 Ferredoxin, critical hydration level, 77 Fish, antifreeze proteins, 151 Flory-Huggins equation, 43 Fluorescence spectroscopy hydration, 84-86 a-lactalbumin and lysozyme, 261-262 Folding, hydration, 142- 143 Food, hydration, 152 Free-energy simulations, hydration, 120- 122 Free interface diffusion, crystallization, 22

G Galactosyltransferase, 250-259 conformational changes, 253-254 forms, 252 interactions a-lactalbumin, 255-25 1 residues, a-lactalbumin and lysozyme, 249 as marker in malignancy, 250-251 metal ion binding, 254-255 molecular weights, 252 occurrence, 250

SUBJECT INDEX

preparation, 251 -252 purification, 251 -252 reactions catalyzed by, 179- 180, 256 relationships of structure to function, 253-255 SH group, 253 stability, 252 substrate structural requirements, 257-258 Glycosides, hydrolysis, 200-20 1 Growth cessation, 19-20

H Hanging-drop technique, crystallization, 30 Heat capacity hydration, 47-50 discontinuities, 132- 133 isotherm, regions, 48-49 a-Helical proteins, water binding to carbony1 groups, 109 Hemoglobin critical hydration level, 77 deoxygenated sickle, osmotic pressuremolar volume isothermals, 58-59 mutation, free-energy simulations, 121 Heterogeneous nucleation, 7- 12 Homogeneous nucleation, 6-7 Hydration, 37-161 ab initio simulations, 120 absorbance spectroscopy, 1 10- 11 1 amide hydrogen exchange, 135 anhydrobiosis, 151 antifreeze proteins, 151- 152 chemistry of, 123-125 transition states, 143- 145 circular dichroism, 111 compressibility, 61 computer simulation accessible surface and thermodynamics, 117-120 molecular dynamics, 112- 1 15 Monte Carlo simulations, 1 15- 117 conformation, 139- 141 coupling parameter, 120 coupling protein and solvent motions, 130-131 critical level, 148- 149

339

denaturation, 52-53 dielectric relaxation, 61 -69 capacitance, 64-66 high frequency, 62-64 KHz and MHz frequencies, 64-67 low frequency, 66, 68 percolation parameter, 66-67 protonic conduction, 64-65 thermal depolarization, 68-69 diffraction, 99- 107 active-site waters, 104- 106 crambin, 102- 103 high-resolution analyses, 106 insulin, 104 lysozyme, 99- 102 myoglobin, 103 dynamics, 134- 136 electron spin resonance, 76-79 electrostatic simulations, 122 empirical valence bond method, 121 end point, 133, 138 enthalpy, 45-46 enzyme activity, 91-95, 135, 144 five-parameter model, 118 fluctuations catalysis and, 148- 149 protein motions and, 148 fluorescence, 84-86 folding, 142- 143 food, 152 force, 56-60 as function of distance between surfaces, 57-58 free-energy simulations, 120- 122 fully hydrated protein protein, 129- 131 solvent, 126- 129 heat capacity, 47-50 discontinuities, 132- 133 heat capacity and spectroscopic properties, 131-134 high-hydration event, 135 hydration shell model, 119 hydrogen exchange, 80-84 infrared and Raman spectroscopy, 107-110 ionization, 50-52 isosteric heat, 45 magnetic susceptibility, 1 12

340

SUBJECT INDEX

Hydration (continued) mass ratio, 43 measurement methods, 38-39 mechanical properties, 98-99 membranes, 149-150 Mossbauer spectroscopy, 88 neutron scattering, 85-87 nonfreezing water, 54-55 diffraction, 55 nuclear magnetic resonance, 54-55 scanning calorimetry, 54 nuclear magnetic resonance, 7 1-76 amount of hydration water, 74-76 nonfreezing water, 54-55 powders, 71-73 solutions, 72-74 percolation model, 69-7 1, 150 percolation theory, 154- 161 perturbation of multilayer water, 79-80 preferential, 5-6 solvation and multicomponent systems, 60-61 process, 38 protein rate processes, 129 protonic conduction and percolation, 145- 146 reverse niicelles, microemulsions, and nonaqueous solvents, 95-96 sorption, 41-45 substrate binding, 146- 147 tetrasaccharide substrate binding, 145 thermodynamic perturbation method, 12 1 time-average properties, 131- 134 200 K transition, 136-137 viscosity, 96-98 volume, 50-51 water networks, 147- 148 Hydration shell, 38,40, 137-139 binding, 139 definition, 138 fluctuations and, 148- 149 folding and, 143 model, 119 properties, 136 thermodynamic properties, 126- 127 time-average properties, 138 Hydrogen exchange amide, 80-82,97-98, 135

hydration, 80-84 lysozyme, 5 1 Hysteresis, sorption, 44-45

I Immunological properties, a-lactalbumin and lysozyme, 272-275 Infrared spectroscopy, hydrated proteins, 107- 110 Insulin, diffraction, 104 Interleukin lp, crystals, 10- 12 Introns, a-lactalbumin and lysozyme, 280-282 Ionization, hydration, 50-52

K K(I), a-lactalbumin binding, 22 1 Kramers theory, modified, 97

L a-Lactalbumin, 174- 176 activity determination, 186- 187, 190- 192 amino acid compositions, 224-227,232 amino acid sequence, 2 11 bovine, 232,238 camel, 240 caprine and ovine, 238-239 comparison with lysozyme, 180- 181, 232-240 equine, 240 guinea pig, 239 human, 239,240-241 rabbit, 239, 288 rat, 240 red-necked wallaby, 240, 287 amino-terminal residues, 247 antigenic determinants, 273-274 apparent heterogeneity, 190 association and aggregation, 267-268 association constants for binding of Ca(II), 297-298 A state, 269-270 basic and acidic groups, 249 calcium-containing crystals, 209 cation binding, 218-222 chain length, 246-247

SUBJECT INDEX

chemical reactivities, 27 1-272 circular dichroism, 263-264 comparison to lysozyme structural elements, 211-212 conformational states in solution, 220-221 crystallographic data, 196- 197 denaturation, 268-27 1 disulfide bridges, 247 early history, 178- 180 electron spin resonance, 265-267 evolution, 276-293 divergence of a-lactalbumin and c-type lysozyme, 286-290 introns and exons, 280-282 mammalian evolution and paleontology, 278-280 models, 286-287,289-290 molecular clocks, 276-278 rapid, 290 fluorescence spectroscopy, 261 -262 fractions, 178- 179 freeze-drying, 295 functions, exclusivity, 290-293 galactosyltransferase activity, 191- 192 genetic variants, 190 helices and P sheets, 2 11- 2 12 immunochemical properties, 272-275 interactions with galactosyltransferase, 255-257 intramolecular distances, 220 invariant residues, 247-248 isolation, 186-187, 190, 296-297 lactose synthase activity, 190- 191, 218-222 ligands for Ca(I1) binding, 248-249 low-angle X-ray scattering, 265 luminescence properties, 26 1 lytic activity, 293 metal ion binding, 216-218 implications for lysozyme, 222-223 monovalent cation effects, 221 neutron diffraction studies, 294 nuclear magnetic resonance, 265-267 occurrence, 186, 188-189,291-292 optical rotary dispersion, 263-265 pH and forms, 2 18 Raman spectroscopy, 262-263 renaturaiion, 268-27 1 . I

34 1

reoxidation, 270-271 residues catalysis, 249 galactosyltransferase interaction, 249 reversible changes at low pH, 187 site-directed mutagenesis, 295 sperm maturation, 298-299 tertiary structure, 269 three-dimensional structure, 293-295 Browne et al. model, 206-207 Ca(I1) binding site, 213 Lewis and Scheraga model, 207-208 models, 206-209 nuclear Overhauser effect studies, 213-214 substrate cleft, 207-208 Warme et al. model, 208-209 water molecules, 211, 213 x-ray crystal structure, 209, 214 transitions, 268 UV absorption spectroscopy, 59 UV difference spectroscopy, 259-261 P-Lactoglobulin, hydrated, thermograms, 52 Lactose marsupial and monotreme milk, 29 1-292 synthesis, 179 Lactose synthase system a-lactalbumin, 2 18-222 galactosyltransferase and a-lactalbumin interactions, 255-257 lysozyme, 292 Langmuir isotherm, 43-44 Lanthanides, a-lactalbumin binding, 2 19 Lipid bilayers, force-distance relationships, 58 Low-angle X-ray scattering, a-lactalbumin and lysozyme, 265 Lysozyme, 174-176 active site, 198-199 activity, 135, 183 amino acid compositions, 224-225, 228-232 amino acid sequence, 274 axis deer stomach mucosa, 242 baboon, 241 bobwhite quail egg white, 244 bovine. 242

342

SUBJECT INDEX

Lysozyme (continued) bovine stomach mucosa, 242 California quail egg white, 244 chachalaca egg white, 246 comparison with a-lactalbumin, 180-181,232-238,240-246 differences in same species, 296 domestic hen egg white, 243 duck egg white, 245-246 echnidna, 287-288 echnidna milk, sequence, 243 equine, 241 guinea hen egg white, 245 phylogenetic analysis, 283, 285 pigeon egg white, 246, 288 pig stomach mucosa, 242 ring-necked pheasant egg white, 245 rodent, 241-242 stomach, 285 turkey egg white, 244-245 amino-terminal residues, 247 antibacterial properties, 298 antigenic determinants, 273-274 apparent specific heat capacity, 48 ArgILys ratios, 284-285 association constants for binding of Ca(II), 297-298 basic and acidic groups, 249, 284-285 cell lysis kinetics, 183- 185 cell lytic action mechanism, 195, 198- 204 catalysis of polysaccharide substrate cleavage, 200-201 cleft, 202 difference electron density study, 198 enzyme-catalyzed hydrolysis, 198, 200 glycoside hydrolysis, 200-20 1 molecular dynamics study, 203 transglycosylation, 201 -202 undistorted ring at site D, 202-203 chain length, 246-247 chemical reactivities, 271-272 circular dichroism, 111, 263-264 comparison to a-lactalbumin structural elements, 211-212 compressibility, 61 cross-reaction, 273 crystallographic data, 196- 197 denaturation, 268-27 1

detection, 185- 186 diffraction, 99- 102 disulfide bridges, 247 early history, 176- 180 echidna, functions, 29 1 enzyme activity, 92-94 ESR spectra, 77-78,265-267 evolution chick-, goose-, phage-, and insecttype, 282-283 divergence of a-lactalbumin and ctype lysozyme, 286-290 introns and exons, 280-282 mammalian evolution and paleontology, 278-280 models, 289-290 molecular clocks, 276-278 stomach, 283-286 fluorescence spectroscopy, 261 -262 freeze-drying, 295 functions, exclusivity, 290-293 genetic engineering, 203 helices and p sheets, 21 1-212 hexasaccharide binding sites, 198,200 'H spin-lattice relaxation, during dehydration, 74-75 hydration dependence of dielectric response, 62-63 hydrogen exchange, 5 1,80-8 1 immunochemical properties, 272-275 implications of a-lactalbumin metal ion binding, 222-223 invariant residues, 247-248 IR spectrum, 107- 108 isolation, 181-184, 295-296 lactose synthase system, 292 ligands for Ca(I1) binding, 248-249 low-angle X-ray scattering, 265 luminescence properties, 261 lytic activity, 292-293 main-chain conformation, 192- 193 as major digestive enzyme in ruminants, 285-286 as marker, 298 metal ion binding, 214-215 molecular chains, 16- 17 molecular dynamics simulation, 112- 113 Monte Carlo simulation, 115- 116 neutron diffraction studies, 294

343

SUBJECT INDEX

neutron scattering, 87 nuclear magnetic resonance, 7 1-73, 265-267 ''0and *H resonances, 75 occurrence, 181 optical rotary dispersion, 263-265 ordered water molecules about, 100- 101 out-exchange of tritium, 81-82 percolation parameters, 66-67 protein-water interactions, 205-206 protonic conduction, 145 Raman spectroscopy, 262-263 relation between crystal size and supersaturation, 18 renaturation, 268-27 1 residues catalysis, 249 galactosyltransferase interaction, 249 side chain hydration, 102 site-directed mutagenesis, 295 site-specific mutagenesis, 274-275 sorption isotherm, 41-42 spin-spin interaction, 140 S-S bond, 270 structures, human and tortoise egg white, 204 tendency to form complexes, 182 tertiary structure, 210 three-dimensional structure, 293-295 types, 181 UV absorption spectroscopy, 59 UV difference spectroscopy, 259-261 water in crystals, 204-206 x-ray crystal structure, 192- 195 Lysozyme-saccharidecomplex, protonic percolation, 70 Lysozyme-watersystem, specific heat, 47

M Magnetic susceptibility, hydration, 112 Malignancy, galactosyltransferase as marker, 250-251 Mammals, evolution and paleontology, 278-280 Membranes, hydration, 149- 150 Metal ion binding a-lactatbumin, 216-218

implications for lysozyme, 222-223 lysozyme, 214-215 Mica, forces between charged surfaces, 56-57 Micelles, reverse, hydration, 95-96 Microemulsion, hydration, 95-96 Microgravity, crystallization in, 26, 29-30 Microheterogeneity, 25 Milk protease, 252 Molecular clocks, a-lactalbumin and lysozyme, 276-278 Molecular preassociation role, in nucleation and crystal growth, 16-17 Monosaccharides, enhanced binding of a-lactalbumin and galactosyltransferase, 257 Monte Carlo simulations free-energy, 121- 122 hydration, 115-117 Multicomponent systems, hydration, 60-61 Mutagenesis site-directed, a-lactalbumin and lysozyme, 295 site-specific,lysozyme, 274-275 Myoglobin diffraction, 103 Mossbauer spectroscopy, 88-90 neutron scattering, 87

N Na(I), a-lactalbumin binding, 22 1 Neon-water system, simulations, 112 Neutral theory of molecular evolution, 277-278 Neutron diffraction studies, a-lactalbumin and lysozyme, 294 Neutron scattering, hydration, 85-87 Nonaqueous solvents, hydration, 95-96 Nuclear magnetic resonance hydration, 71-76 a-lactalbumin and lysozyme, 265-26 nonfreezing water, 54-55 Nucleation competition with crystal growth, 17- 9 conditions for, 22 determination of conditions, 12- 13 heterogeneous, 7- 12

344

SUBJECT INDEX

Nucleation (continued) homogeneous, 6-7 rate, 7-8, 12

0 Optical rotary dispersion, a-lactalbumin and lysozyme, 263-265 Osmotic pressure-molar volume isothermals, deoxygenated sickle hemoglobin, 58-59 Ovalbumin, denaturation, 53 Oxidation, cysteine, 24 Oxymyoglobin, neutron diffraction analysis, 103

P Pancreatic trypsin inhibitor molecular dynamics simulation, 113 Monte Carlo simulation, 115, 117 Parvalbumin, molecular dynamics simulation, 113- 114 Percolation, protonic conduction and, 145- 146 Percolation model, 41, 69-71, 150 Percolation theory, 128- 129, 154- 161 conductivity, 159 critical threshold, 154 critical volume fraction, 156- 157 dependence on lattice type, 157-158 exponents for two and three dimensions, 157, 159 finite-size effects, 160- 161 invariant quantities, 156- 159 percolation probability, 155, 159, 161 percolation threshold, 155, 157 phenomena modeled by, 155-156 Percolation transition, 70 Phosphatidylcholine bilayers, force-distance relationships, 58 Phosphorescence, hydration, 84-85 Protein motions, fluctuations and, 148 Protein rate processes, hydration, 129 Protein-water bond, 7 1 Protein-water interactions, lysozyme, 205-206 Protonic conduction hydration dependence, 64-65 percolation and, 145- 146

Protonic percolation, 70 Pseudomonas indigofera, isocitrate lyase crystals, 9- 10 Purity, crystallization, 23-25 Purple membrane critical exponent for protonic percolation, 66-67 dielectric measurements, 150 percolation parameters, 66-67

R Raman spectroscopy hydrated proteins, 107- 110 a-lactalbumin and lysozyme, 262-263 Rayleigh scattering, Mossbauer radiation, 88,90 Renaturation, a-lactalbumin and lysozyme, 268-271 Reoxidation, a-lactalbumin, 270-27 1 Ribonuclease, circular dichroism spectra, 111 Ribonuclease A, enthalpy dependence on water content, 46 Rough surface model, 13-14

S Scanning calorimetry, nonfreezing water, 54 Screw dislocation model, 14 Seeding, crystallization, 23 SH group, galactosyltransferase, 253 Solution turbulence, crystallization, 29 Solvation, preferential, hydration, 60-61 Solvent-accessiblesurface, simulations, 117-120 Sorption hydration, 41-45 isotherm, 75 lysozyme, 41-42 Sperm maturation, a-lactalbumin, 298-299 Spin-lattice relaxation rate, versus concentration, 74-75 Spin-spin interaction, lysozyme, 140 S-S bond, lysozyme, 270 Streptavidin crystals, 8-9 Streptomyes avidinii, streptavidin crystals, 8-9

345

SUBJECT INDEX

Substrate binding, hydration, 146- 147 Successive automated grid search method, 26-28 Supersaturation, 3-4 dependence of nucleation and growth rates, 18 nucleation rates, 7-8 Surface kinetics, control of crystal growth, 13-14 Surface motion, 129-130 Surface nucleation model, 14 Surface water crystallographic estimate, 127- 128 dynamic properties, 128

T Tb(II), a-lactalbumin binding, 218-219 Temperature shift, crystallization, 21 -22 TEMPONE correlation time, 78-79 ESR spectra, 77-78 Tetrasaccharide substrate, binding, 145 Thermal depolarization, hydration, 68-69 Thermodynamic perturbation method, 121 Thermodynamics hydration, simulations, 117- 120 transfer of solvent into interface, 126- 127 Transfer free energies, cyclohexane, 1 18 Transglycosylation, 20 1-202 Transition states, chemistry, hydration and, 143-145

Transport phenomena in protein crystal growth, 15- 16 rate, control of crystal growth, 13

U UDP-glucose, as donor substrate, 257-258 Unfolding, hydration contribution, 119 Urea, enzyme activity, 94 Urease, enzyme activity, 94 UV absorption spectroscopy, a-lactalbumin and lysozyme, 259 UV difference spectroscopy, a-lactalbumin and lysozyme, 259-261

V Vapor diffusion, crystallization, 21 Viscosity, hydration, 96-98 Volume, hydration, 50-5 1

W Water nonfreezing, hydration, 54-56 networks, hydration, 147- 148

Y Young’s modulus, temperature-dependent change, 98

Z Zn(II), a-lactalbumin binding, 219-220

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