Advances in Protein Chemistry Volume 41

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ADVANCES IN PROTEIN CHEMISTRY Volume 41

This Page Intentionally Left Blank

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

San Diego New York Boston London Sydney Tokyo Toronto

This book is printed on acid-free paper. @

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.

ACADEMIC PRESS, INC. San Diego, California 92101 United Kingdom Edition published by ACADEMIC PRESS LIMITED 24-28 Oval Road, London NW17DX

LIBRARY OF CONGRESS CATALOG CARD NUMBER: 44-8853

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

PRINTED IN THE UNITED STA'IES OF AMERICA 91

92 93 94

9 8 7 6 5 4 3 2 1

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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1 3 3 6 13 17 19 20 23 25 26 32 33

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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38 41 61 99 112

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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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SUBJECT 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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3 3 6 6 7 7

12 13 13 13 14 15 16 17 19 20 20 20 21 21 22 22 23 23 25 26 26 30 32 33

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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PATRICIA C. WEBER

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.

PHYSICAL PRINCIPLES OF PROTEIN CRYSTALLIZATION

3

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

4

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-


5

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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PATRICIA C. WEBER

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


7

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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PATRICIA C . WEBER

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


13

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,


15

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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PATRICIA C. WEBER

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-


17

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).


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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PATRICIA C. WEBER

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-


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-

22

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.


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

24

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-


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.


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.


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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PATRICIA C. WEBER

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.


33

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.


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.

38

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


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


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)


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


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.


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


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).


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


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).


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


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


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


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


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


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


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


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


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


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


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


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


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


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


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


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


-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


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


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


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


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


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.


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


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


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


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


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


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


- *

- 0.4

I

k 0

-

0

9-

-

I

*

0 t-

*L

-0.3 0

a LL

tA

- 0.2

I

*

-

z

'4

@

c

-0 I

L

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


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.


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


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


-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


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


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

L3 2.2

f

2.8

-

2.1

2.7

-

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2.6

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2.5

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1.8

2.4

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(-5)

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2.3 2.2

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FIG.24. The average lifetime of a-chymotrypsinogen A as a function of water content (h). Left and right ordinate scales are for different methods of averaging the two components detected in analysis of the decay curves. From Fucaloro and Forster (1985).

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.


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


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


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

h 7

' v

x 0

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temperature (K) FIG. 26. Temperature dependence of various properties of myoglobin crystals: 0 ,frequency of the 0-D band maximum (IR); ---, dielectric relaxation time of water (schematic); , Lamb-Mossbauer factor, f o , after subtracting the harmonic mode (schematic). From Doster et al. (1986).

90


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

1

00.2 .31

"d

0

:€

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).


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


0

0.2

0.6

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


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).

94


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-


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,,

96

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


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


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


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


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


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 ( &(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 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).

190

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

192

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.


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. 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


209

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

210

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


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).


213

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


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

215

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


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-


217

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


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


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


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


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


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


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


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

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


299

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


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


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.


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


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


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.


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.


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


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.


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


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


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.


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.


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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