Fibrous Proteins: Amyloids, Prions and Beta Proteins, Volume 73 (Advances in Protein Chemistry)

CONTENTS b-Structures in Fibrous Proteins Andrey V. Kajava, John M. Squire, and David A. D. Parry I. Introduction . ...

Author: John M. Squire | David A.D. Parry | Andrey Kajava

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CONTENTS

b-Structures in Fibrous Proteins Andrey V. Kajava, John M. Squire, and David A. D. Parry

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Characteristics of Simple b-Structures. . . . . . . . . . . . . . . . . . . . . III. Diversity of b-Structural Fibrous Folds Revealed by Crystallographic Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Recent Advances in Structural Studies of Amyloid and Prion Fibrils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 4 7 10 12 13

b-Silks: Enhancing and Controlling Aggregation Cedric Dicko, John M. Kenney, and Fritz Vollrath

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. b-Silk: An Optimized System for Controlled Assembly and Aggregation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Role and Function of b-Sheet Assembly in Silk Proteins . . . . . IV. Fibril Assembly: Amyloid Nature of Silk? . . . . . . . . . . . . . . . . . . V. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

18 22 30 39 41 42

b-Rolls, b-Helices, and Other b-Solenoid Proteins Andrey V. Kajava and Alasdair C. Steven

I. II. III. IV. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Diversity and Classification of b-Solenoids . . . . . . . . . . . . . . . . . Capping and Bulging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Multistranded b-Solenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Relationship Between b-Solenoid Structures and Their Amino Acid Sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v

56 61 70 71 74

vi

CONTENTS

VI. Relationship Between b-Solenoid Structures and Their Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Evolution of b-Solenoid Proteins . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

85 87 89 90

Natural Triple b-Stranded Fibrous Folds Anna Mitraki, Katerina Papanikolopoulou, and Mark J. van Raaij

I. II. III. IV.

Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Crystal Structures of Viral Fibers. . . . . . . . . . . . . . . . . . . . . . . . . . Stability, Folding, and Assembly of Fibrous Proteins. . . . . . . . . Future Research Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

98 99 111 116 118

Structure, Function, and Amyloidogenesis of Fungal Prions: Filament Polymorphism and Prion Variants Ulrich Baxa, Todd Cassese, Andrey V. Kajava, and Alasdair C. Steven

I. II. III. IV. V. VI. VII. VIII. IX.

Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Prion Domains and Functional Domains . . . . . . . . . . . . . . . . . . Filament Formation In Vivo and In Vitro . . . . . . . . . . . . . . . . . . . Filament Formation and Prion Conversion Are Based on Amyloidosis of the Prion Domains . . . . . . . . . . . . . . . Experimentally Derived Constraints on Prion Filament Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structural Models for Prion Amyloid Filaments. . . . . . . . . . . . . Other Structural Considerations. . . . . . . . . . . . . . . . . . . . . . . . . . Prion Variants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

127 135 137 143 151 157 162 166 171 172

X-Ray Fiber and Powder Diffraction of PrP Prion Peptides Hideyo Inouye and Daniel A. Kirschner

I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Prion Hypothesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

182 186

CONTENTS

III. IV. V. VI. VII. VIII.

Sequence Analysis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Prion Alanine-Rich Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Amyloidogenic Core Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . Polyalanine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Polyglutamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

vii 189 191 198 199 203 205 206

From the Polymorphism of Amyloid Fibrils to Their Assembly Mechanism and Cytotoxicity Laurent Kreplak and Ueli Aebi

I. II. III. IV. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Polymorphism of Amyloid Fibrils. . . . . . . . . . . . . . . . . . . . . . . . . Soluble Forms of Amyloid Peptides. . . . . . . . . . . . . . . . . . . . . . . Depicting Intermediate Stages of Amyloid Fibril Assembly . . . What Is the Mechanism of Small Oligomer-Induced Cytotoxicity?. . . . . . . . . . . . . . . . . . . . . . . . . . VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

217 219 223 223 226 229 229

Structural Models of Amyloid-like Fibrils Rebecca Nelson and David Eisenberg

I. II. III. IV. V. VI.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Refolding Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gain-of-Interaction Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Models of Natively Disordered Proteins . . . . . . . . . . . . . . . . . . . Fibril Properties and Their Relation to Structural Models . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

236 239 243 257 264 271 272

AUTHOR INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

283 309

SUBJECT INDEX. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

b‐STRUCTURES IN FIBROUS PROTEINS By ANDREY V. KAJAVA,* JOHN M. SQUIRE,{ AND DAVID A. D. PARRY{ *Centre de Recherches de Biochimie Macromolećulaire, CNRS FRE‐2593, 1919 Route de Mende, 34293 Montpellier Cedex 5, France; { Biological Structure and Function Section, Biomedical Sciences Division, Imperial College London, London SW7 2AZ, United Kingdom; { Institute of Fundamental Sciences, Massey University, Palmerston North, New Zealand

I. II. III. IV. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Characteristics of Simple b‐Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Diversity of b‐Structural Fibrous Folds Revealed by Crystallographic Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Recent Advances in Structural Studies of Amyloid and Prion Fibrils . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 4 7 10 12 13

Abstract The b‐form of protein folding, one of the earliest protein structures to be defined, was originally observed in studies of silks. It was then seen in early studies of synthetic polypeptides and, of course, is now known to be present in a variety of guises as an essential component of globular protein structures. However, in the last decade or so it has become clear that the b‐conformation of chains is present not only in many of the amyloid structures associated with, for example, Alzheimer’s Disease, but also in the prion structures associated with the spongiform encephalopathies. Furthermore, X‐ray crystallography studies have revealed the high incidence of the b‐fibrous proteins among virulence factors of pathogenic bacteria and viruses. Here we describe the basic forms of the b‐fold, summarize the many different new forms of b‐structural fibrous arrangements that have been discovered, and review advances in structural studies of amyloid and prion fibrils. These and other issues are described in detail in later chapters.

I.

Introduction

Elucidation of the three‐dimensional structures of b‐structural fibrous proteins has attracted the interest of scientists for more than 50 years. In the early days, the objects of these studies were predominantly the naturally occurring fibrous assemblies obtained from b‐silk and stretched mammalian ADVANCES IN PROTEIN CHEMISTRY, Vol. 73 DOI: 10.1016/S0065-3233(06)73001-7

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Copyright 2006, Elsevier Inc. All rights reserved. 0065-3233/06 $35.00

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Fig. 1. The basic arrangements of b‐strands in hydrogen‐bonded b‐sheets (A) parallel chains, (B) antiparallel chains. Green spheres of different sizes denote side chain groups

FIBROUS b‐STRUCTURES

3

b‐keratin (Astbury and Street, 1931), but the crystalline structures formed by some synthetic polypeptides (Fraser and MacRae, 1973) were also investigated in detail. An important outcome of these studies was a description of the two basic b‐structural arrangements found in proteins: the parallel and antiparallel pleated b‐sheet structures (Fraser et al., 1969; Pauling and Corey, 1951; Fig. 1A and B). Most of these b‐sheet structures are nonplanar (i.e., twisted), as shown initially by Fraser et al. (1971) for feather keratin, but subsequently seen widely in virtually all crystalline globular proteins (Salemme and Weatherford, 1981). At the same time, significant progress was achieved in establishing the orientation of the b‐crystallites that composed the pleated sheet structures in b‐silk, b‐keratin, and the other fibrous polypeptide structures (Bradbury et al., 1960; Fraser and MacRae, 1973). More recently, research on fibrous b‐proteins has been stimulated by the observation that amyloids, prion fibrils, and a variety of denaturated globular proteins have cross‐b structures (Fig. 1C and D), in which the polypeptide chains are oriented perpendicular to the plane of the fibrils axis (Blake and Serpell, 1996; Caughey et al., 1991; Eanes and Glenner, 1968; Kirschner et al., 1986). The incidence of amyloid fibrils in important human diseases has attracted considerable efforts to solve their structures at the atomic level. Despite this, however, the structure of the amyloid fibril, and in particular the lateral packing of the b‐strands and their orientation (parallel vs antiparallel) within the b‐sheets, remains unknown. This failure may be attributed in part to the fact that methods of determining high‐resolution structure (protein crystallography and NMR spectroscopy) cannot be used because of the polymeric character and insolubility of the fibrils involved. Accordingly, X‐ray fiber diffraction, electron microscopy (EM), optical spectroscopy, and other biophysical approaches have been the principal sources of data underlying the models of b‐structural fibrils presented to date. Over the last 13 years, there have been major advances in the study of fibrous b‐proteins. In particular, this period has been marked by a rapid emergence of new structural information. First, a number of crystal structures having elongated b‐structural fibrous topologies have been resolved by X‐ray crystallography, thanks to improved expression and crystallization strategies. Second, several new experimental techniques, including solid‐ state NMR, scanning transmission EM mass measurements, and electron directed either toward (large spheres) or away from (small spheres) from the reader. Hydrogen bonds are shown by red dotted lines. Other colors follow the standard CPK scheme. (C) Chain folding back onto itself in a cross‐b sheet. (D) Stacking of several sheets as in (C); the spacing of the stacks, shown as 11 A˚, is actually very variable depending on the nature of the R groups.

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KAJAVA ET AL.

paramagnetic resonance spectroscopy of spin‐labeled derivatives, have been applied successfully and these have provided significant constraints on the structural models for b‐silk, amyloid, and prion fibrils. One of the aims in preparing this book has been to provide an overview of the progress made in the elucidation of the b‐fibrous proteins over the past decade.

II.

Characteristics of Simple b‐Structures

To set the scene for the detailed chapters in the rest of the book, we describe here the main features of the simple b‐structural components of proteins. The parallel and antiparallel b‐structures in Fig. 1A and B show certain characteristic features. There is a repeat along the chain direction (vertical in Fig. 1A and B) which consists of two‐amino acid residues and is often about 6.5‐ to 7‐A˚ long. The b‐sheet is not planar but pleated to permit the side chains (R groups) of the amino acids to project out from the plane of the backbone‐pleated sheet. In Fig. 1A and B, the backbone‐pleated sheets are in the plane of the page and the R groups are imagined projecting above and below this plane. The other relatively constant dimension in b‐structures is the repeat distance in the direction of the hydrogen bonding between adjacent chains. In the antiparallel b‐structures this distance is about 9.6 A˚, but this contains two chains so there is a marked repeat at half of this, about 4.8 A˚. In the silks (Dicko et al., this volume) and in stretched keratin, the chain axis is normally parallel to the fiber axis direction, as envisaged in Fig. 1A and B for a ‘‘vertical’’ fiber axis. However, early studies of synthetic polypeptides (Bradbury et al., 1960) showed that some structures existed where the chain axis was perpendicular to the direction of stroking or stretching when the polypeptide solutions were oriented before drying. This was termed the cross‐b structure (Fig. 1C); it was also found to exist in a number of denatured globular and fibrous proteins (see summary in Fraser and MacRae, 1973). The term ‘‘cross‐b structure’’ was originally used to imply an antiparallel arrangement of b‐strands lying perpendicular to the fibril axis. It is now used more generally, however, to describe any chain arrangement of b‐strands (parallel, antiparallel, or mixed) with a chain orientation perpendicular to the fibril axis. The in‐plane spacings in cross‐b structures are much the same as in Fig. 1A and B, except that the hydrogen‐bonded direction (9.6‐ and 4.8‐A˚ repeats) is now along the fiber axis and the in‐chain repeat of 7 A˚ is perpendicular to the fiber axis. For b‐crystallites in general (i.e., for those structural elements in which the chain directions lie either in a similar direction to the fibril axis or which lie approximately perpendicular to it), the repeat in the third


5

dimension has proved to be quite variable. The hydrogen‐bonded sheets depicted in Fig. 1A–C can sometimes stack together as shown in Fig. 1D for a cross‐b antiparallel sheet. The exact separation of the sheets is much more variable than the other repeats and depends crucially on the nature of the side chains (R groups) of the amino acids in the sheets and how they pack together. This intersheet spacing has been observed to be as low as 5 or 6 A˚ in some simple synthetic polypeptides, and can vary between about 8 and 16 A˚ in some prion structures (see Inouye and Kirschner, this volume), although the crystallographic repeat in this direction can be multiples of this basic intersheet spacing depending on precisely how the sheets are arranged. In many amyloid structures this third direction, if it involves sheet packing (see later), often has a repeat around 10–11 A˚. There are several different techniques with which to categorize b‐structures into those with their chain axis oriented along the fiber axis or those which are cross‐b types, but among the most powerful is the method of fiber diffraction (usually with X‐rays, but sometimes using neutrons or electrons). Figure 2 illustrates schematically the differences that might be observed in the fiber diffraction pattern of well‐aligned samples depending on whether the chains are oriented along (A, B) or perpendicular to (C, D) the fiber axis. Once again the 7‐A˚ repeat along the chain (call it C for Chain) and the 9.6‐A˚ repeat (or half of this) in the hydrogen‐bonded direction (call it H for Hydrogen‐bonded) are relatively constant features, whereas the peaks associated with the intersheet distance (call it S for Sheet) can have a variety of positions and strengths. The C‐repeat along the chain is roughly the pitch of a 2/1 helix of amino acids where the subunit repeat is 3.5 A˚, so the 3.5‐A˚ repeat gives a meridional peak in the diffraction pattern from the parallel b‐structure (Fig. 2B), with the 7‐A˚ repeat showing up as off‐meridional intensity on the first layer‐line. The H and S directions are perpendicular to this, so for the axially aligned b‐structures they show up as intensities along the equator of the pattern. Once again, in the antiparallel sheet the 9.6‐A˚ H direction corresponds to the separation of two chains, so there is a strong pseudo‐repeat after 4.8 A˚ which often makes this peak much stronger than the 9.6‐A˚ peak (which in fact may not be observed). In the case of diffraction from the cross‐b structure, the repeat in the fiber axis direction is the hydrogen‐bonded H‐repeat of 9.6 or 4.8 A˚. The S‐repeat still shows up on the equator, but the C‐repeat along the chain axis has now switched from the meridian in Fig. 2C to the equator in Fig. 2D. Finally, a number of observed amyloid diffraction patterns, which are often quite disoriented, giving arced diffraction peaks, show simply the interchain spacing (H) at around 4.8 A˚ on the meridian and the intersheet spacing at around 10–11 A˚ on the equator. In Figs. 1C, D, and 2C, the

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Fig. 2. The differences that might be observed in the fiber diffraction patterns from oriented samples of antiparallel b‐structures depending on whether the chains are aligned along (A, B) or perpendicular to (C, D) the fiber axis. Color coding on (A, B) as in Fig. 1.

antiparallel chains are shown folding back on themselves so that a single sheet can be formed from just one chain. However, many well‐studied b‐structures, some natural and some synthetic, are aggregates of very short peptides, some of which are not long enough to fold back on themselves even once. In these cases, there is often ambiguity about whether adjacent chains are parallel or antiparallel and the intersheet stacking can be very variable. As discovered recently in structures solved by protein crystallography, there is in fact a great diversity of chain organizations in naturally occurring b‐structural folds, as summarized in the next section.


III.

7

Diversity of b‐Structural Fibrous Folds Revealed by Crystallographic Studies

During the past few years, a new set of b‐structural fibrous folds has emerged. One of these, the parallel b‐helix, was first described for bacterial pectate lyase in 1993 (Yoder et al., 1993). Since then more than a hundred crystal structures with similar or related fibrous morphologies have been solved. These have revealed a great diversity of b‐structural folds that can be categorized into at least five distinct groups (Fig. 3). They include b‐solenoids (Fig. 3A), triple‐stranded b‐solenoids (Fig. 3B), triangular cross‐b prisms (Fig. 3C), triple b‐spirals (Fig. 3D), and spiral b‐hairpin stacks (staircases; Fig. 3E). The crystal structures generally have axial dimensions that are comparable to their lateral ones and are, therefore, in a sense, not strictly fibrous. However, these structures are built of axially stacked repetitive structural blocks. This arrangement, in principle, allows ready elongation to form fibrils by the simple addition of recurrent blocks. The other common property of the fibrous b‐proteins is the repetitive character of their amino acid sequences. The majority of the known b‐fibrous proteins are located on the surfaces of either bacteria or viruses. A significant portion of these proteins forms homotrimers. The most frequently occurring b‐fibrous folds are based on solenoidal windings of the polypeptide chain. Each coil in the solenoid has an axial rise of about 4.8 A˚ and corresponding b‐strands in successive coils align to form parallel b‐sheets (Fig. 3A). The number of the known b‐solenoid proteins, which include b‐helices and b‐rolls, is now large enough to support their detailed analysis and classification. Kajava and Steven (this volume) present a systematic account of these structures distinguished by their handedness, twist, oligomerization state, and coil shape. This survey has also revealed some relationships between the amino acid sequences of b‐solenoids and their structures and functions. This has implications for structural prediction of other b‐solenoids and for elucidation of amyloid fibril structures. A recently discovered subset of triple‐stranded b‐helices from bacteriophage tail proteins (alternatively termed ‘‘triple‐stranded b‐solenoids’’) represents another distinct group of b‐fibrous folds (Fig. 3B). In these structures, three identical chains related by threefold rotational symmetry wind around a common axis. These chains form unusual parallel b‐sheets with no intra‐ and only intermolecular b‐structural hydrogen bonding. Kajava and Steven (this volume) survey the distinguishing structural features of the known triple‐stranded b‐solenoids, also documenting their notable diversity and differences in comparison to the single‐stranded b‐solenoids.

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KAJAVA ET AL.

A

B

b -Solenoid

C

Triple-stranded b -solenoid

D

Cross-b -prism E

Triple b -spiral

Spiral b -hairpin staircase

Fig. 3. Representative structures of five principal groups of b‐structural fibrous folds that have recently been established by X‐ray crystallographic studies. Arrows denote b‐strands. b‐Strands that belong to the same chains have the same color.

The abundance, location, stability, and folding of the triple‐stranded b‐helices are also reviewed in a chapter by Mitraki, Papanikolopoulou, and van Raaij, which is dedicated to triple b‐stranded fibrous folds in the viral fibers. Mitraki and colleagues also overview the other distinctive family of b‐fibrous folds, called the triple b‐spirals (Fig. 3D). The b‐spiral folds are more complicated than the solenoidal fold, with long central b‐strands that hold the trimer together through interchain hydrogen bonds, and


9

interactions of the apolar side chains and short peripheral b‐strands to stabilize the structure. The distinctive structural property of this trimeric fold is that its longest core b‐strands run parallel to the fiber axis in contrast to most of the other b‐fibrous folds representing variations of the cross‐b structure. Mitraki and coauthors also describe the structures of b‐structural globular domains identified in the viral fibers and demonstrate their crucial role in the correct folding of the adjacent b‐fibrous folds such as triple‐stranded b‐helices and b‐spirals. Finally, the chapter by Mitraki et al. illustrates some crystallographic strategies that may lead to the discovery of even more new b‐fibrous modules from the trimeric fibers of viruses. For completeness of classification, it is pertinent to mention two other families of the b‐fibrous folds. One of these is a homotrimeric structure that resembles a triangular prism with equivalent sides formed by antiparallel cross‐b sheets (Fig. 3C). The internal side of these b‐sheets is composed of apolar side chains, while the opposite side consists primarily of polar side chains. This fold was found in the crystal structure of autotransporter protein Hia from Haemophilus influenzae (Yeo et al., 2004) and in the tailspike protein of Salmonella typhimurium phage P22 (Schuler et al., 2000). The other distinctive b‐fibrous fold was discovered among the surface proteins of pathogenic Gram‐positive bacteria (Fernandez‐Tornero et al., 2001; Ho et al., 2005) or their bacteriophages (Hermoso et al., 2003). This is a single‐stranded b‐fibrous fold with b‐hairpins as repetitive structural units (Fig. 3E). The b‐hairpins extend perpendicularly from the axis and the relation between adjacent hairpins can be approximated by a threefold screw‐axis transformation characterized by a 90–120 unit rotation and an axial translation of about 10 A˚, thereby creating a left‐handed superhelix. This fold resembles a spiral staircase with b‐hairpins as the steps. The ‘‘spiral b‐hairpin staircase’’ fold was found in choline‐binding domains in the pneumococcal virulence factor LytA (Fernandez‐Tornero et al., 2001) and endolysin from pneumococcal bacteriophage Cp‐1 (Hermoso et al., 2003) as well as in carbohydrate‐binding domain of toxin A from Clostridium difficile (Ho et al., 2005). It is worth mentioning that, in contrast to the other b‐fibrous folds, the fibrils generated by the spiral b‐hairpin staircase fold can readily curve due to the absence of hydrogen bonding between adjacent b‐hairpins. Thus, the classical pleated b‐sheet structures have now been supplemented by several new b‐structural fibrous folds that have been established by X‐ray crystallographic studies. As a consequence, today, the b‐fibrous folds represent a more diverse class of fibrous structures than those defined by the a‐ or collagen‐helices.

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KAJAVA ET AL.

Recent Advances in Structural Studies of Amyloid and Prion Fibrils

Over the past decade, significant progress has been made in understanding the structural arrangements of prion and amyloid fibrils. This can be attributed to the establishment of several b‐solenoid folds that are consistent with the available constraints imposed by the structure of amyloid fibrils. Furthermore, new experimental techniques such as solid‐ state NMR, scanning transmission EM mass measurements, and electron paramagnetic resonance spectroscopy of spin‐labeled derivatives have provided a number of new constraints for modeling amyloid and prion structures. An important achievement of this work was the establishment of parallel and in‐register arrangement of b‐strands in several amyloid fibrils. These included b‐amyloid, a‐synuclein, human amylin, and yeast Ure2p (Benzinger et al., 1998; Chan et al., 2005; Der‐Sarkissian et al., 2003; Jayasinghe and Langen, 2004). Based on this and other experimental data, several new structural models for amyloid and prion fibrils with parallel in‐register stacking of b‐strands have been formulated (Govaerts et al., 2004; Guo et al., 2004; Kajava et al., 2004, 2005; Petkova et al., 2002; Ritter et al., 2005; Wang et al., 2005; Fig. 4). These results have effectively put an end to the dominance of those models characterized by antiparallel b‐sheet arrangements. It has also revealed that the parallel b‐sheets in these prion and amyloid fibrils differ from the antiparallel ones observed in the fibrils formed by short (7–10 residue) fragments of the same peptides (Balbach et al., 2000; Griffiths et al., 1995). These data clearly indicate the possibility that the unconstrained short peptides may not have the same structure/ properties as they do in the context of a full‐length peptide. Fibrils of the first mammalian prion protein discovered, namely PrP (Prusiner, 1991), proved to be difficult to study as a consequence of poor in vitro reproduction of what were homogeneous well‐ordered fibrils in vivo (Baskakov and Bocharova, 2005). Nevertheless, a considerable body of data has been collected on the structure and formation of these fibrils. In this volume, Kirschner and Inouye review current knowledge of PrP prion pathologies and summarize X‐ray fiber and powder diffraction studies on the N‐terminal fragments of prion proteins. They also compare structures of PrP peptide assemblies with those of the PrP‐related polyalanine and polyglutamine peptides. Recently, prions have been found in fungal systems (Wickner, 1994) and this has advanced the field considerably, due to the improved experimental tractability of these prions. Baxa and colleagues (this volume) have focused on the structures of the fungal prion fibrils and in so doing have both summarized current experimental constraints and appraised the various models proposed. The authors have concluded

11


A

B

b-Helical models

C

b-Superpleated model

Fig. 4. New structural models for amyloid and prion filaments with the parallel and in‐register arrangement of b‐strands in the b‐sheets. b‐Strands are denoted by arrows. The filaments are formed by hydrogen‐bonded stacks of repetitive units. Axial projections of single repetitive units corresponding to each model are shown on the top. Lateral views of the overall structures are on the bottom. (A) The core of a b‐helical model of the b‐amyloid protofilament (Petkova et al., 2002). Two such protofilaments coil around one another to form a b‐amyloid fibril. (B) The core of a b‐helical model of the HET‐s prion fibril (Ritter et al., 2005). The repetitive unit consists of two b‐helical coils. (C) The core of a superpleated b‐structural model suggested for yeast prion Ure2p protofilaments and other amyloids (Kajava et al., 2004).

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KAJAVA ET AL.

that two parallel b‐structural folds, the superpleated b‐structure (Kajava et al., 2004) and a specific b‐helical formulation (Ritter et al., 2005), are the most valid candidates to explain the known experimental data on Ure2p and HET‐s prion filaments. The polymorphism of the yeast prion fibrils and its possible structural basis have also been discussed in several chapters of this volume. Polymorphism appears to be a common property of prion and amyloid fibrils. The same amyloidogenic peptide can form different fibril structures depending on slight changes in the fibrillogenesis conditions. This adds complexity to what is already a difficult problem in analyzing the structure of amyloid fibrils. Kreplak and Aebi (this volume) have surveyed the polymorphism of fibrils formed by b‐amyloid peptides, calcitonin, human amylin, and other proteins as observed by EM. They have linked this phenomenon with differences in the structure of small intermediate oligomers that initiate fibrillogeneses. Approaches that allow a study of such oligomers and provide information on the relation of the intermediates with the toxicity of the amyloid peptides have also been discussed. Nelson and Eisenberg (this volume) have provided a detailed review of the current structural models of prion and amyloid fibrils. Despite the fact that atomic‐level structures of amyloid‐like fibrils have yet to be determined, many models of these fibrils have been proposed. Nelson and Eisenberg categorize these models into three classes: (i) Refolding models, in which the protein has different structures in the native and fibrillar states; (ii) Gain‐of‐Interaction models, which propose a largely native‐like structure for proteins in the fibril; and (iii) Natively Disordered models, which are formed by peptides whose native state is not structured. It was shown that the cores of several models contain a packing of the b‐strands similar to that in the so‐called cross‐b spine structure. This has recently been determined at atomic resolution using X‐ray diffraction of the crystal formed from a seven‐residue peptide from the yeast prion Sup35 (Nelson et al., 2005).

V. Conclusions The contributions to this volume demonstrate that structural studies of fibrous b‐proteins, as well as prion and amyloid fibrils, have advanced rapidly thanks in large part to improved experimental techniques and better theoretical analysis of the ever‐increasing structural data. It is also possible to learn from studies of naturally occurring silks (Dicko et al., this volume) how variations in the conditions of production of silk threads from the same protein can produce a variety of b‐structures with very distinct


13

properties. Further progress in studies of b‐structures is likely to lead to the discovery of new b‐fibrous folds, to the determination of more precise structural models for amyloid‐like fibrils, and to a better understanding of the factors that determine why they fold as they do. Since diseases like Alzheimer’s are likely to affect a gradually increasing proportion of the population as life expectancy increases, this is not an unimportant task. The observed abundance of the b‐fibrous folds among virulence factors of bacteria and viruses also indicates that the fibrous b‐proteins will be an attractive target for future structural studies, especially in the context of emerging infectious threats.

References Astbury, W. T., and Street, A. (1931). X‐ray studies of the structures of hair, wool and related fibres. I. General. Trans. R. Soc. Lond. A230, 75–101. Balbach, J. J., Ishii, Y., Antzutkin, O. N., Leapman, R. D., Rizzo, N. W., Dyda, F., Reed, J., and Tycko, R. (2000). Amyloid fibril formation by A beta 16–22, a seven‐residue fragment of the Alzheimer’s beta‐amyloid peptide, and structural characterization by solid state NMR. Biochemistry 39, 13748–13759. Baskakov, I. V., and Bocharova, O. V. (2005). In vitro conversion of mammalian prion protein into amyloid fibrils displays unusual features. Biochemistry 44, 2339–2348. Benzinger, T. L., Gregory, D. M., Burkoth, T. S., Miller‐Auer, H., Lynn, D. G., Botto, R. E., and Meredith, S. C. (1998). Propagating structure of Alzheimer’s beta‐ amyloid(10–35) is parallel beta‐sheet with residues in exact register. Proc. Natl. Acad. Sci. USA 95, 13407–13412. Blake, C., and Serpell, L. (1996). Synchrotron X‐ray studies suggest that the core of the transthyretin amyloid fibril is a continuous beta‐sheet helix. Structure 4, 989–998. Bradbury, E. M., Brown, L., Downie, A. R., Elliott, A., Fraser, R. D. B., Hanby, W. E., and Macdonald, T. R. R. (1960). The ‘‘cross‐beta’’ structure in polypeptides of low molecular weight. J. Mol. Biol. 2, 276–281. Caughey, B. W., Dong, A., Bhat, K. S., Ernst, D., Hayes, S. F., and Caughey, W. S. (1991). Secondary structure analysis of the scrapie‐associated protein PrP 27‐30 in water by infrared spectroscopy. Biochemistry 30, 7672–7680. Chan, J. C., Oyler, N. A., Yau, W. M., and Tycko, R. (2005). Parallel beta‐sheets and polar zippers in amyloid fibrils formed by residues 10‐39 of the yeast prion protein Ure2p. Biochemistry 44, 10669–10680. Der‐Sarkissian, A., Jao, C. C., Chen, J., and Langen, R. (2003). Structural organization of alpha‐synuclein fibrils studied by site‐directed spin labeling. J. Biol. Chem. 278, 37530–37535. Eanes, E. D., and Glenner, G. G. (1968). X‐ray diffraction studies on amyloid filaments. J. Histochem. Cytochem. 16, 673–677. Fernandez‐Tornero, C., Lopez, R., Garcia, E., Gimenez‐Gallego, G., and Romero, A. (2001). A novel solenoid fold in the cell wall anchoring domain of the pneumococcal virulence factor LytA. Nat. Struct. Biol. 8, 1020–1024. Fraser, R. D. B., and MacRae, T. P. (1973). ‘‘Conformation in Fibrous Proteins and Related Synthetic Polypeptides.’’ Academic Press, London, New York.

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Fraser, R. D. B., MacRae, T. P., Parry, D. A. D., and Suzuki, E. (1969). Structure of beta‐ keratin. Polymer 10, 810–826. Fraser, R. D. B., MacRae, T. P., Parry, D. A. D., and Suzuki, E. (1971). The structure of feather keratin. Polymer 12, 35–56. Govaerts, C., Wille, H., Prusiner, S. B., and Cohen, F. E. (2004). Evidence for assembly of prions with left‐handed beta‐helices into trimers. Proc. Natl. Acad. Sci. USA 101, 8342–8347. Griffiths, J. M., Ashburn, T. T., Auger, M., Costa, P. R., Griffin, R. G., and Lansbury, P. T. (1995). Rotational resonance solid‐state NMR elucidates a structural model of pancreatic amyloid. J. Am. Chem. Soc. 117, 3539–3546. Guo, J. T., Wetzel, R., and Xu, Y. (2004). Molecular modeling of the core of Abeta amyloid fibrils. Proteins 57, 357–364. Hermoso, J. A., Monterroso, B., Albert, A., Galan, B., Ahrazem, O., Garcia, P., Martinez‐ Ripoll, M., Garcia, J. L., and Menendez, M. (2003). Structural basis for selective recognition of pneumococcal cell wall by modular endolysin from phage Cp‐1. Structure 11, 1239–1249. Ho, J. G., Greco, A., Rupnik, M., and Ng, K. K. (2005). Crystal structure of receptor‐ binding C‐terminal repeats from Clostridium difficile toxin A. Proc. Natl. Acad. Sci. USA 102, 18373–18378. Jayasinghe, S. A., and Langen, R. (2004). Identifying structural features of fibrillar islet amyloid polypeptide using site‐directed spin labeling. J. Biol. Chem. 279, 48420–48425. Kajava, A. V., Baxa, U., Wickner, R. B., and Steven, A. C. (2004). A model for Ure2p prion filaments and other amyloids: The parallel superpleated beta‐structure. Proc. Natl. Acad. Sci. USA 101, 7885–7890. Kajava, A. V., Aebi, U., and Steven, A. C. (2005). The parallel superpleated beta‐structure as a model for amyloid fibrils of human amylin. J. Mol. Biol. 348, 247–252. Kirschner, D. A., Abraham, C., and Selkoe, D. J. (1986). X‐ray diffraction from intraneuronal paired helical filaments and extraneuronal amyloid fibers in Alzheimer disease indicates cross‐beta conformation. Proc. Natl. Acad. Sci. USA 83, 503–507. Nelson, R., Sawaya, M. R., Balbirnie, M., Madsen, A. O., Riekel, C., Grothe, R., and Eisenberg, D. (2005). Structure of the cross‐beta spine of amyloid‐like fibrils. Nature 435, 773–778. Pauling, L., and Corey, R. B. (1951). Configurations of polypeptide chains with favored orientations around single bonds: Two new pleated sheets. Proc. Natl. Acad. Sci. USA 37, 729–740. Petkova, A. T., Ishii, Y., Balbach, J. J., Antzutkin, O. N., Leapman, R. D., Delaglio, F., and Tycko, R. (2002). A structural model for Alzheimer’s beta‐amyloid fibrils based on experimental constraints from solid state NMR. Proc. Natl. Acad. Sci. USA 99, 16742–16747. Prusiner, S. B. (1991). Molecular biology of prion diseases. Science 252, 1515–1522. Ritter, C., Maddelein, M. L., Siemer, A. B., Luhrs, T., Ernst, M., Meier, B. H., Saupe, S. J., and Riek, R. (2005). Correlation of structural elements and infectivity of the HET‐s prion. Nature 435, 844–848. Salemme, F. R., and Weatherford, D. W. (1981). Conformational and geometrical properties of beta‐sheets in proteins. I. Parallel beta‐sheets. J. Mol. Biol. 146, 101–117. Schuler, B., Furst, F., Osterroth, F., Steinbacher, S., Huber, R., and Seckler, R. (2000). Plasticity and steric strain in a parallel beta‐helix: Rational mutations in the P22 tailspike protein. Proteins 39, 89–101.


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Wang, J., Gulich, S., Bradford, C., Ramirez‐Alvarado, M., and Regan, L. (2005). A twisted four‐sheeted model for an amyloid fibril. Structure 13, 1279–1288. Wickner, R. B. (1994). [URE3] as an altered URE2 protein: Evidence for a prion analog in Saccharomyces cerevisiae. Science 264, 566–569. Yeo, H. J., Cotter, S. E., Laarmann, S., Juehne, T., St Geme, J. W., III, and Waksman, G. (2004). Structural basis for host recognition by the Haemophilus influenzae Hia autotransporter. EMBO J 23, 1245–1256. Yoder, M. D., Keen, N. T., and Jurnak, F. (1993). New domain motif: The structure of pectate lyase C, a secreted plant virulence factor. Science 260, 1503–1507.

b‐SILKS: ENHANCING AND CONTROLLING AGGREGATION By CEDRIC DICKO,* JOHN M. KENNEY,{ AND FRITZ VOLLRATH* {

*Zoology Department, Oxford University, OX1 3PS, United Kingdom; Physics Department, East Carolina University, Greenville, North Carolina 27858

I.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Structure and Function Relationship in Silks. . . . . . . . . . . . . . . . . . . . . . . . . . B. Apparent Paradox of ‘‘Hydrophobic Proteins’’ and ‘‘Water Spinning’’ ... C. Aims of This Review. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. b‐Silk: An Optimized System for Controlled Assembly and Aggregation . . . . A. Model Assembly in Silks and Enrichment for b‐Structures . . . . . . . . . . . . . B. Silk Proteins Stability and Solubility in Solution: Sol–Gel Transition . . . III. Role and Function of b‐Sheet Assembly in Silk Proteins . . . . . . . . . . . . . . . . . . . A. Monitoring and Studying Silk Protein Behavior . . . . . . . . . . . . . . . . . . . . . . . B. b‐Transition: The Trademark of Silk Proteins . . . . . . . . . . . . . . . . . . . . . . . . . C. Factors Governing the Transition to b‐Structures. . . . . . . . . . . . . . . . . . . . . . D. Irregular Sequence Units and Heterogeneous Structures . . . . . . . . . . . . . . IV. Fibril Assembly: Amyloid Nature of Silk? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Spinning Versus Growth: Length and Time Scale Change . . . . . . . . . . . . . B. Missing Cross‐b Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

18 18 21 21 22 22 25 30 30 31 35 37 39 39 40 41 42

Abstract It appears that fiber‐forming proteins are not an exclusive group but that, with appropriate conditions, many proteins can potentially aggregate and form fibrils; though only certain proteins, for example, amyloids and silks, do so under normal physiological conditions. Even so, this suggests a ubiquitous aggregation mechanism in which the protein environment is at least as important as the sequence. An ideal model system in which forced and natural aggregation has been observed is silk. Silks have evolved specifically to readily form insoluble ordered structures with a wide range of structural functionality. The animal, be it silkworm or spider, will produce, store, and transport high molecular weight proteins in a complex environment to eventually allow formation of silk fibers with a variety of mechanical properties. Here we review fiber formation and its prerequisites, and discuss the mechanism by which the animal facilitates and modulates silk assembly to achieve controlled protein aggregation.

ADVANCES IN PROTEIN CHEMISTRY, Vol. 73 DOI: 10.1016/S0065-3233(06)73002-9

17


18

DICKO ET AL.

I. Introduction A.

Structure and Function Relationship in Silks

Silks have evolved to be some of nature’s most impressive composite materials (Craig, 1997). Silk fibers (Denny, 1980) and glues (Vollrath et al., 1990) are not only one of the toughest polymers known, but they have a number of other characteristics that, although less well known, make them an interesting as well as an important object for research in the general areas of biopolymers (Vollrath and Knight, 2004), protein folding, biomimetics and the coevolution of behavior, morphology, and function (Vollrath, 2000a). Silk proteins (spidroins in spiders and fibroins in Lepidoptera insects) are assembled into well‐defined nanofibrillar architectures (Craig and Riekel, 2002; Eby et al., 1999; Inoue et al., 2000b, 2001; Li et al., 1994; Putthanarat et al., 2000; Vollrath et al., 1996). Spidroins and fibroins are largely constructed from two chemically distinct repetitive motifs or ‘‘blocks’’ (Table I), an insoluble crystalline block and a soluble less‐crystalline block (Craig, 2003; Fedic et al., 2002; Hayashi and Lewis, 2000; Hayashi et al., 1999). The crystalline blocks are composed of short side‐chained amino acids in highly repetitive sequences that give rise to b‐sheet structures. Remarkably, the overall mechanical properties (Denny, 1980; Gosline et al., 1999a; Vollrath, 2000b) of silks seem to depend less on the complex hierarchical organization of silk on a microscopic level (Inoue et al., 2000a; Knight and Vollrath, 2002; Vollrath, 1999) and more on the overall molecular structure (Craig, 2003; Fedic et al., 2003; Gosline et al., 2002; Sponner et al., 2005a) which by a complex spinning process (Liu et al., 2005; Vollrath and Knight, 2001) is formed into a material where nanoscale interactions predominate (Porter et al., 2005; Vollrath and Porter, 2006). From the early X‐ray diffraction studies (Lucas et al., 1960; Marsh et al., 1955; Rudall and Kenchington, 1971b; Warwicker, 1954) to the more modern spectroscopic results (Beek et al., 2000, 2002; Grubb and Jelinski, 1997; Riekel et al., 1999b; Sirichaisit et al., 2003), the identification of the different conformations present in silk fibers and their role in determining silks’ mechanical properties have highlighted the presence of three major types of silks: (i) the rarely observed a‐silk (Hepburn et al., 1979), (ii) the cross‐b silks (Geddes et al., 1968), and (iii) the commonly observed collinear (i.e., with strands parallel to the fiber axis) b‐silks (Craig, 1997; Rudall and Kenchington, 1971a). Interestingly, a‐silks and cross‐b silks can undergo, on postprocessing (i.e., stretching), a transition to the collinear b‐structure. The structural and sequence studies have highlighted two interesting prerequisites to fiber formation. First, the animal must control the size and

Table I Structure Function Relationship in Silk Proteins

Silk type MA

Proteins (ratio?)a

Function Dragline radial threads

MASp 1 and 2 (?)

Predicted structure from sequenceb (GA)n/(A)n b‐structure

Amino acid (%)c Gly (38), Ala (29), SSC (70), PC (21)

GPGGX/GPGQQ ‘‘b‐spiral’’ GGX 31 helix n¼2–8 Auxiliary threads

MISp 1 and 2 (?)

FLAG

Sticky spiral threads Cocoon silks

FLAG

Acinous

Coating, prey wrapping

AcSp1

Pyriform

Attachment disk

?

CYL

TuSp1

(GA)n/(A)n b‐structure GGX 31 helix n¼2–8 spacer GPGGX ‘‘b‐spiral’’ GGX 31 helix spacer (A)n, (S)n, (SA)n, (SQ)n, GX, n¼0–3 No specific repeats but 200 amino acids iteration along sequence ?

Gly (40), Ala (35), SSC (80), PC (19)

Gly (36), Pro (14), SSC (51), PC (27) Ser (20), Ala (27), SSC (56), PC (45) Gly (8), Ala (9), Ser (5), Pro (5), SSC (22), PC (50) Gly (10), Ala (11), Ser (10), Pro (10), SSC (31), PC (52)

Fiber degree of crystallinity (%)f

Fiber extensibility (%)g

b‐Sheet

15–30

35

High concentration: helical/molten globule Helix‐like

b‐Sheet

?

>35

b‐Spiral

b‐Turns

None

>200

Helix‐like

b‐Sheet and b‐turn

?

25

Helix‐like


?

80

Helix‐like


?

?

Low concentration: disorder/PPII

19

(continued)

b‐SILKS

MI

Conformational change in solutione

Structure in solutiond

TABLE I (continued ) Predicted structure from sequenceb

Structure in solutiond

(GAGAGS)5–15 Heavy, b‐sheet (GX)5–15 light chains fibroins and b‐turns/helices P25 (6–6‐1) GAAS spacer

Gly (43), Ala (30), Ser (12), SSC (85), PC (23)

Antheraea Cocoon pernyi

Heavy chain fibroin

(S1–2A11–13), GX1–4 GGX, GGGX

Galleria Cocoon mellonella

Heavy, light chains fibroins and P25

(S1–2A1–4)1–2, GLGGLG, XGGXG GPX spacer

Gly (27), Ala (43), Ser (11), SSC (81), PC (26) Gly (28), Ala (22), Ser (17), SSC (67), PC (26)

Low‐concentration H‐fibroin: disorder/PPII/b‐ turn type II silkI) High‐concentration helical Disorder and helical structures ?

Silk type Bombyx mori

a

Function Cocoon

Conformational change in solutione

Fiber degree of crystallinity (%)f

Fiber extensibility (%)g

b‐Sheet (silk II)

40–50

10–20

b‐Sheet

40

35

b‐Sheet

?

80

Proteins. Sequences and predicted secondary structures: MA, MI, FLAG (Hayashi et al., 1999), CYL (Garb and Hayashi, 2005), Tubuliform (Tian and Lewis, 2005), Aciniform (Hayashi et al., 2004), Bombyx mori (Inoue et al., 2000a; Zhou et al., 2000), Galleria mellonella (Zurovec and Sehnal, 2002), Antheraea pernyi (Sezutsu and Yukuhiro, 2000). c Amino acid composition: MA, MI, FLAG, and CYL from Dicko et al. (2004b), Acinous and Pyriform from unpublished data, Bombyx mori, Antheraea pernyi, and Galleria mellonella from sequences. d Structure in solution: Bombyx mori (Iizuka and Yang, 1966; Yao et al., 2004), MA, MI, FLAG, and CYL (Dicko et al., 2004b), Acinous and Pyriform (Fig. 8), Antheraea pernyi (Tsukada et al., 1994). The helix‐like structure is loosely defined as a structure with a CD spectrum similar to myoglobin. b‐Spiral structure is defined as a super helical structure formed of ‘‘straight’’ sections and b‐turns. e Conformational changes under denaturing conditions (Fig. 8) and fiber formation. f Degree of crystallinity: MA (Riekel et al., 1999a), Bombyx mori, and Antheraea pernyi (Iizuka, 1965). g Fibers extensibility: MA (Vollrath, 1999), MI (Vollrath unpublished), FLAG (Gosline et al., 1999a,b), CYL (Dicko et al., 2004b), Acinous (Hayashi et al., 2004), Bombyx mori, Antheraea pernyi, Galleria mellonella (Denny, 1980). MA, major ampullate; MI, minor ampullate; FLAG, flagelliform; CYL, cylindriform; PPII, polyproline II. Small side chains (SSC)¼glycineþalanineþserine, Polar chains (PC)¼aspartic acidþthreonineþserineþglutamic acidþtyrosineþlysine þhistidineþarginine. b

DICKO ET AL.

Amino acid (%)c

20

Proteins (ratio?)a

b‐SILKS

21

concentration of b‐crystals (Iizuka, 1965; Urs et al., 1993) to be able to extrude the silk without accidental aggregation and achieve good mechanical properties. Second, silk proteins (despite their large size and overall hydrophobicity) are processed in an aqueous environment at ambient temperature and pressure. This suggests an extremely tight control of conformation, solubility, and transport of the protein(s) along the spinning pathway.

B. Apparent Paradox of ‘‘Hydrophobic Proteins’’ and ‘‘Water Spinning’’ Before exploring the fine details of silk construction, we may take a look at some structural properties of the virtually insoluble fiber as found in nature. For example, both in Bombyx mori (a moth bred uniquely for silk production) cocoon silk and in Nephila (a wild orb‐weaving spider) safety‐line silk, selection pressure on silk structure and function have favored the high crystallinity (Table I) and the high molecular weight needed to achieve exceptional mechanical properties (Denny, 1980; Gosline et al., 1999b; Vollrath, 2000b). These two fibers, however, although both virtually insoluble, behave differently in the presence of water: Bombyx fibers will only be weakly plasticized whereas Nephila dragline shows a dramatic and reversible water uptake yielding fiber shrinkage or supercontraction (Grubb and Ji, 1999; Work, 1981; Yang et al., 2000). The ability to plasticize is reflected by the extensibility of the fibers (Table I) and highlights the presence of mobile ‘‘soluble’’ structures within the silks (Beek et al., 1999; Sapede et al., 2005). Another important aspect is the effect of water on the prespun silk proteins. At the beginning of the process of fiber formation, in the secretory glands, the silk proteins are exceptionally soluble in water (20–30% w/v). This is unexpected given the hydrophobic and repetitive nature of silk proteins (Fig. 1). In a recent review, Bini et al. (2004) explore and propose how silk genes have favored such an arrangement. Specific amphiphilic bloc constructs (Fig. 1) promote appropriate solubility, mesogenic units for liquid crystal formation and controllable assembly (Foo et al., 2006). The key to the process is a progressive enrichment in b‐structures without premature aggregation, thus allowing macroscopic structure and order to develop as well as local conformational rearrangement to take place. The mechanism, however, by which soluble hydrophobic proteins are controlled and processed into mechanically robust fibers remains unclear.

C. Aims of This Review To understand the role of silk protein design and b‐structure assembly, it is important to consider the exact sequence of events in the aggregation

22

DICKO ET AL.

Fig. 1. Hydrophobic plot of (A) B. mori heavy chain fibroin (P05790 b.) and (B) Nephila clavipes spidroin 1 (P19837 f.). The hydropathy is plotted as a function of residue position along the gene sequences, each residue’s hydropathy is weighted using the hydropathy of its 13 nearest neighbor residues. The figure illustrates the apparent paradox of an aqueous spinning of hydrophobic silk proteins. To overcome the high hydrophobicity in B. mori, the silkworm produces a transportable and processable protein complex (Fig. 2), whereas with spidroin, the amphiphilic nature of sequence gives enough flexibility to allow water (hydrogen bonds) and hydrophobic interactions to operate and fold the protein (Fig. 2).

and orientation of silk protein molecules in the natural extrusion process. After all, it is here where the development of a macroscopic order accompanied by local conformational transition is effected. Here we review the molecular mechanisms that might be behind the generation of ‘‘stable’’ and processable silk proteins and examine how those mechanisms may have been optimized in spiders and insects. For the purpose of this review, we focus on the implications of our insights into silk form and function to elucidate amyloid structure and formation, although we will extend the discussion in order to include fiber‐forming proteins in general.

II.

‐Silk: An Optimized System for Controlled Assembly and Aggregation A. Model Assembly in Silks and Enrichment for b‐Structures

1.

Fiber Construction: Matching and Aligning the Sequences

It is now clear from the study of silk fiber formation in lepidoptera and spiders (Akai, 1998; Iizuka, 1966; Kerkam et al., 1991; Knight and

b‐SILKS

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Vollrath, 2002; Magoshi et al., 1994; Viney, 1997; Viney et al., 1994; Vollrath and Knight, 2001) that the construction of the silk fiber core (Sehnal and Zurovec, 2004; Sezutsu and Yukuhiro, 2000; Zurovec and Sehnal, 2002) involves a precise locking and docking of the chains (Fig. 2) and domains identified in Table I. The accuracy by which the chains will fold and align with opposite segments will determine the crystallinity and resistance to swelling or water plasticization of the fiber. The variations in sequences, for example in the three lepidoptera shown in Table I, are reflected in the final silk mechanical properties. One interesting aspect of the folding process is the transformation of apparently disordered prespun silk proteins into b‐sheet‐rich structures. Specifically, we must ask how the sequence design promotes proteins crystallization and lamellar liquid crystal orientation (Knight and Vollrath, 1999b, 2002; Oroudjev et al., 2002). The answer appears to lie in the amount and size of crystals required to achieve a specific strength of the fiber (Bini et al., 2004; Sehnal and Zurovec, 2004). Hence, much depends on how well the chains are aligned during processing.

2. b‐Sheet Enrichment The silk fiber is formed near the end of the duct during a dramatic phase transition resulting from rapid flow elongation in a draw down taper within the spinning duct (Knight et al., 2000; Vollrath and Knight, 2001). The linear velocity of the protein through the duct increases exponentially before the draw down taper, suggesting that wall shear may play a part in the transition to solid silk. A controlled flow elongation as well as water extraction provides a progressive increase in birefringence correlated to an increase in b‐sheet structure in the duct (Knight et al., 2000; Rossle et al., 2004; Viney et al., 1994). In a recent study, Jin and Kaplan (2003) demonstrate the formation of silk fibroin aggregates in the presence of polyethylene glycol, and present a step by step model for fiber formation based on the principle of micelle formation, and driven by dehydration as well as flow elongation. During this process, hydrophobic chains are exposed to the solvent, but because of the molecules’ high free energy, water solvation is unfavorable and phase separation followed by aggregation predominates. Other additional phenomena may contribute to the progressive enrichment in b‐sheet structure. For example, silk glands produce multiple proteins and the interactions of these proteins facilitate fiber formation and contribute to the size and amounts of crystallinity (Craig, 2003; Lee, 2004; Sehnal and Zurovec, 2004; Sponner et al., 2005b). Furthermore,

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Fig. 2. Model assembly in B. mori and Nephila dragline silks. Left: Illustration of an assembly model and the intracellular transport of the elementary unit of silk fibroin. In this model, a nascent heavy‐chain fibroin (H‐chain) is synthesized on the membrane‐bound polysome. In the endoplasmic reticulum (ER), the H‐chain is covalently linked to the light‐chain fibroin (L‐chain) via a disulfide bond. The H–L dimers thus formed, together with an N‐glycosylated P25, are then assembled into the H6L6P251 complex. The main driving forces for the complex assembly are hydrophobic interactions between P25 and the H‐chains and hydrogen bonding between the P25’s N‐linked oligosaccharide chains and the H‐chains. The elementary unit is then transported to the Golgi apparatus for further posttranslational modifications. Finally, the elementary unit is excreted into the posterior part of the silk gland lumen. The silk protein assembly unit ensures solubility during transport until fiber formation. Modified and redrawn from Inoue et al. (2000a), Zhou et al. (2000). (With permission of Journal of Biological Chemistry. Copyright of the American Society for Biochemistry and Molecular Biology, Inc.) Right: Illustration of an assembly model and the extracellular transport of spider silk major ampullate (MA) protein in orb web spiders. The process occurs via a lamellar liquid crystalline assembly into nanofibrils to form the final silk thread. A diagrammatic optical section of the secretory part (A‐ and B‐zones) of the gland and the spinning duct shows how silk fibers are formed. Initially the silk protein units are secreted and excreted as a tight hexa‐columnar packed arrangement of amphiphilic rod‐shaped molecules. In the early stage of fiber formation (in the duct), a flow elongation field will progressively unwind and align the silk molecules to form nanofibrils. The final assembly of the nanofibrils takes place at the draw down taper, where the solid fiber is eventually formed. The lumen of the gland has been represented as much wider in proportion to length with only a small number of bilayer discs (top left) and the epithelium has been shown on only one side of the duct and gland. The dotted lines represent the molecular director field. This lies at right angles to the slow axis of polarization as a result of the assembly of the compactly wound, rod‐ shaped molecules of spidroin into bilayered discs of the nematic discotic phase. These are present as an escaped nematic texture in the gland and first half of the duct. Modified and redrawn from Knight and Vollrath (2002).

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posttranslational modifications have been identified, and most likely these modulate protein solubility (Bini et al., 2004). In summary, the formation of silk fibers involves superstructures such as, possibly, micelles and/or molecular rods (Akai, 1998; Jin and Kaplan, 2003; Knight and Vollrath, 2002) that are dependent on the packing and conformation of the individual protein units. The relationship, however, between the shapes of these superstructures and the various forms of protein conformation remains elusive (Valluzzi and Jin, 2004). Seeking to clarify this issue we will examine, in Section II.B, the role of shape and extended network formation modulating solubility, stability, and assembly.

B. Silk Proteins Stability and Solubility in Solution: Sol–Gel Transition Spidroins and fibroins share several features with amyloids and prions including low complexity and nonrandomness of the amino acid sequence (Kenney et al., 2002; Wise, 2001; Wootton, 1994). The silks are predominantly composed of small side chain residues, alanine, glycine, and serine (Table I; Craig et al., 1999; Lucas et al., 1960). The ratio of these small side chains, however, varies with species and silk function (Craig et al., 1999; Fedic et al., 2002). The natural evolution of spider silk with a predominance of small side chains suggests the need for greater conformational flexibility with less selectivity for specific secondary structures. From a biological point of view, this provides the animal with a silk fiber precursor capable of providing a wider range of possible spinning regimes. In this context, B. mori silk (compared to wild silkworm, such as Antheraea pernyi) has apparently been inbred so much (Asakura and Kaplan, 1993) that the formation of the fiber requires a rigorous sequence of events with only a limited amount of space for error. Nevertheless, despite our rapidly increasing knowledge about silk protein sequences, very little is known about the part played by gene design (Tatham and Shewry, 2000) in the stability and solubility of the prespun silk and in the final property of silk fibers across a wider variety of spiders and insects.

1. b‐Structure: Silk Most Stable Form Spiders and insects both excrete prepacked silk proteins (Inoue et al., 2000a; Knight and Vollrath, 1999a) from the glandular silk cells into the saclike storage glands, which may occur in a large variety of shapes and sizes (Akai, 1998; Kovoor, 1987; Tillinghast and Townley, 1994; Vollrath and Knight, 2004). In these glands, the silk proteins are stored in a stable soluble state prior to fiber formation. The conversion to an insoluble silk

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Fig. 3. Solubility of silk proteins in solution as a function of time. Low solubility corresponds to protein aggregation. The fast and slow aggregations are observed in vitro (Dicko et al., 2004a), whereas the stable helical conformation (storage structure) is observed in vivo (Dicko et al., 2004b,d). This illustrates the inherent instability of silk protein in solution and shows the b‐sheet polymorph structure as the most stable form. In other words, the spiders actively control and modulate the unavoidable silk protein aggregation prior to fiber formation.

fiber occurs in the duct after exiting the gland. However, the silk feedstock in the gland is only stable under specific conditions (Asakura et al., 1993) as is apparent in the dissection of the dragline glands where, if done without care, extracting the silk proteins will result in accelerated aggregation (Fig. 3). Moreover, it is remarkable that the silk proteins with an apparent high solubility in the gland will readily precipitate to form predominantly disordered aggregates (Dicko et al., 2004c, 2005; Liu et al., 1998). Phase diagram studies of fibroin (Magoshi et al., 1994; Sohn et al., 2004) suggest that disordered silk states are drawn to the more stable b‐sheet structures in solution. Figure 4 shows similar phenomena in spider dragline silk in solution (Dicko et al., 2004c). With time and temperature, the silks are progressively phase separating and precipitating. Interestingly, the rate of change is slower at 5 C compared to 20 C, which suggests a hydrophobically driven change. This behavior suggests that prespun silk is stored in a metastable state and that the animal achieves stability by controlling the kinetic rate at which the proteins phase separate and aggregate. A possible mechanism for such tight control is illustrated in Fig. 5. Clearly, increasing the protein concentration has a dramatic impact on the secondary structures of silk proteins in solution. The low concentration silk protein solution at 1% w/v is dominated by disordered structures or equally possible a polyproline II type structure (Sreerama and Woody, 2003).

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Fig. 4. Time‐induced conformational change of spider silk protein (spidroin) in solution. Solutions of silk proteins at 1% w/v in distilled water were monitored using circular dichroism. The graph shows a change in secondary structure with time. The silk proteins underwent a kinetically driven transition from a partially unfolded structure to a b‐sheet‐rich structure (from Dicko et al., 2004c). (□) after 0 days, (○) after 1 day, and (Δ) after 2 days. The conformational change appeared faster at 20 C compared to 5 C, suggesting a hydrophobically driven mechanism. (Copyright 2004 American Chemical Society.)

As the concentration increases, the circular dichroism spectra show more and more helical structures (Dicko et al., 2004c). This concentration effect in Nephila major ampullate (dragline) silk shows a cooperative folding to a molten type globule structure whereby the intermediate, comparable to a molten globule (Baldwin, 1991; Ptitsyn, 1995), could provide compactness (Viney et al., 1994) without rigid packing of secondary structure. This results in substantial mobility of the side chains as well as of larger parts or domains of the proteins (Baldwin, 1991; Dobson, 1992; Uversky et al., 1997). This mechanism would provide the spider with a structurally complex, storable and transportable protein ready to undergo a transition to a solid fiber. Individually, the silk molecules will adopt a shape and conformation dictated by the competition to hydrate polar and nonpolar moieties (Hossain et al., 2003; Jin and Kaplan, 2003). In addition, to achieve fiber formation and optimal axial stiffness, the system must organize and lock the molecules in their extended configuration (Donald and Windle, 1992)

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Fig. 5. The effect of protein–protein interactions on Nephila edulis major ampullate circular dichroism spectra in solution. A change in secondary structure with increasing concentration is observed. At low concentration (minimal protein–protein interactions) silk proteins appear partially unfolded in solution. At higher concentration (higher protein–protein interactions) silk proteins refold into a helix‐like structure, most likely a molten‐like globule (from Dicko et al., 2004c). This final molten structure would facilitate local chain rearrangement while preserving the global structure for protein storage and transport. (Copyright 2004 American Chemical Society.)

and at the same time control the amount of cross‐linking to prevent high viscosity. But, unlike a‐helical structures, b‐strands are not stable as isolated secondary structures (Nesloney and Kelly, 1996) and will gain stability by engaging in side‐to‐side interaction with another strand. Thus, the sheet formed can now associate laterally or in a face‐to‐face manner allowing the construction of an intermolecular network. One can speculate on the b‐network and what has previously been identified as a gel or hydrogel state during silk storage (Akai, 1998; Hossain et al., 2003; Ochi et al., 2002b). Interestingly, rheological studies (Terry et al., 2004) have shown a reversible sol–gel transition on acidification, which in turn suggests that (in readiness to be spun) the proteins stored in the gel state are transformed to a soluble state before being further processed. NMR data (Asakura et al., 1983; Hronska et al., 2004), however, show no compelling evidence for a b‐sheet structured gel state in the glands. Several studies (Asakura et al., 2001; Dicko et al., 2005; Heslot, 1998; Lazo and Downing, 1999; Valluzzi et al., 2002) have focused their effort toward the understanding of silk stability and to describing the secondary structure population present in the prespun silk. One can, however, hypothesize that the formation of the gel state involves the competition between

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Fig. 6. Structural stability of major ampullate silk protein in constrained Nephila edulis. The graph shows a time series of circular dichroism spectra of major ampullate (MA) protein at 1% w/v in distilled water. The spiders prior to dissection were prevented from spinning, but fed and watered for at least 2 weeks. With time, the secondary structure of silk protein is becoming more and more ‘‘disordered.’’ The arrow indicates increasing time (days). Note that the amino acid composition of the silk protein was similar to that of a native N. edulis spider. Interestingly, silk protein extracted from the constrained spider did not respond to denaturing conditions (detergents, alcohols, pH, and salts; Dicko et al., 2004a, 2005).

intramolecular metastable b‐sheet structures and more stable extended structures linked to irreversible aggregation. In this context, noteworthy is an experiment conducted on constrained spiders (Dicko et al., 2004a, 2005). Figure 6 shows for spiders ‘‘force‐fed’’ without being allowed to spin silk, how the conformation of the proteins in solution is drawn toward the disordered state. The proteins started a transition to a ‘‘random coil’’ structure over time—rather than b‐sheet structure formation. Interestingly, the amino acid composition of the silk proteins of these constrained spiders was similar to those of free‐range spiders. Furthermore, the solutions were soluble even at high concentration for a long period of time and did not respond to any denaturants known to induce b‐sheet formation (Dicko et al., 2004a, 2005). Such behavior would support the idea of a predominantly intramolecular interaction and collapse of the chains to form disordered or natively unfolded proteins (Uversky, 2002). In summary, the dynamics of the hierarchical interaction between silk proteins in solution suggest that spiders and insects are trading long range crystallinity for local conformational transitions, thus allowing the

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formation of a spatiotemporal pattern (Turing, 1952) in the form of a gel state. This suggests that chain flexibility and extensibility to control propagation of the ‘‘silk pattern’’ may simply be controlled by nanoscale interactions (Porter et al., 2005; Vollrath and Porter, 2006) and departure from a random network type polymer to an elastomeric network (Urry et al., 2002). Turing’s famous paradigm (1952) ‘‘short‐range activation coupled to long ‐range inhibition is a natural recipe for spatiotemporal pattern formation’’ seems to be the most appropriate way to describe the complex arrangement and processing of silk proteins. In Section III, we will explore how spatial and temporal control of silk protein geometry and chemistry play a role in b‐sheet assembly.

III.

Role and Function of ‐Sheet Assembly in Silk Proteins A.

Monitoring and Studying Silk Protein Behavior

In the study of silk proteins, three main problems are limiting the amount of information available. First, silk protein structures are highly dependent on the preparation method with the direct consequence of extreme sensitivity to conformational change and aggregation. Second, it is difficult to obtain good quality‐oriented structures particularly with silk I, which explain the wealth of information on the insoluble silk II phase found in fibers compared to scarce complete data on the precursor protein. Third, is the difficulty in assessing the protein content and the individual contributions to observable parameters such as a CD spectrum. Because of these limitations, characterization of silk proteins has mainly consisted of understanding the solid fiber structure, while the prespun silk structures remain a mystery (Table I). Studies on B. mori prespun silk have identified two main polymorphs: silk II (b‐sheet structure found in the solid fiber) and silk I which is thought to be dominated by short‐range and/or ‘‘disordered’’ structures (Asakura et al., 1985, 1994). Similar results were found in spiders (Dicko et al., 2004c; Hijirida et al., 1996; Hronska et al., 2004; Kenney et al., 2002). In addition to traditional X‐ray techniques to study silk (Bram et al., 1997; Lotz and Cesari, 1979; Riekel et al., 1999a; Warwicker, 1960), other structural tools have helped unravel various aspects of silk protein conformation. These include solid‐state NMR (Asakura et al., 1983, 1988, 1994; Beek et al., 2000, 2002) studies of native and regenerated silk together with and studies of isotopically edited silks, which have dramatically improved the model of structure distribution within silk fibers (Beek et al., 2000, 2002).

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Optical spectroscopy, on the other hand, has contributed to dynamic studies on silk fibers (Monti et al., 1998; Rousseau et al., 2004; Young et al., 2002), films (Chen et al., 2001, 2002; Minoura et al., 1990; Sonoyama et al., 1997; Tsukada et al., 1994), and dilute solutions (Canetti et al., 1989; Iizuka and Yang, 1966, 1968; Yang et al., 2004), but have had limited success when applied to highly concentrated in vivo like solutions of prespun silk. One technique that has emerged recently is the state‐of‐the‐art technique synchrotron radiation‐based circular dichroism (SRCD) with the potential to give structural information and dynamic data on silk proteins (Dicko et al., 2004c; Wallace, 2000; Wallace and Janes, 2001). We will discuss in Section III.B, the impact of SRCD on understanding the assembly and dynamics of silk protein behavior at near in vivo conditions. Another, promising avenue to understand silk protein conformation and assembly is the use of model peptides. Although not recent (Fraser and MacRae, 1973; Lotz et al., 1974), studies of silk‐based peptide from chemical synthesis, DNA recombinant technology, and computer simulation (Anderson et al., 1994; Asakura et al., 2003; Fahnestock et al., 2000; Fossey et al., 1991; Heslot, 1998; Kaplan, 1998; Wilson et al., 2000) have shown that selected repeats of silk proteins can be transformable hydrogels, elastomers, or regular thermoplastics and that with a proper design they can function as diverse molecular machines (Altman et al., 2003; Heslot, 1998; Kaplan, 1998; Urry, 1998).

B. b‐Transition: The Trademark of Silk Proteins Spiders and insects spinning silks are examples of the success of the polymer principle for optimal axial stiffness: ‘‘the molecules must not only be straight and aligned with the tensile axis, but must also be in their most extended conformation with the backbone bond rotation angles set to give the longest chain possible’’ (Donald and Windle, 1992; Lemstra et al., 1986). But in the case of synthetic polymers this optimal stiffness will require high pressure and temperature, whereas in natural silks it happens in water at ambient temperature and atmospheric pressure (Vollrath and Knight, 2001). The key appears to be in a judicious use of b‐sheet structures and their appropriate control, thus preventing high viscosity and catastrophic aggregation. The b‐transition is a key feature and trademark in silks (Craig, 2003), whether the final product (Fig. 7) will be a high‐performance fiber or will have other functions (Craig, 1997). The role, functionality, and diversity of each silk raise the interesting question whether selection pressures on the final fiber properties are mirrored (at, ultimately, the molecular level) in the precursor liquid proteins. Figure 8 shows SRCD spectra of six of the

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Fig. 7. Silk gland distribution from a typical orb weaver spider (modified from Vollrath, 2000b). Within its abdomen the spider produces up to seven different silks in different glands, all with specific functions (copyright 2000, Elsevier).

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Fig. 8. Temperature‐induced conformational changes in major and minor ampullate (MA and MI), flagelliform, cylindriform, pyriform, and acinous silk proteins. The graphs show the circular dichroism spectra of the various silk proteins at 20% w/v in distilled water at 20 C (black curve) and 80 C (gray curve). The initial solutions at 20 C were progressively heated and left to equilibrate for 15 min at every 5 C step prior to collecting a CD spectrum. The increase in temperature gave a progressive and irreversible change in protein structure (data not shown). All silks produced a recognizable b‐sheet/b‐turn‐rich structure instead of a traditionally expected disordered structure at high temperature. The mid‐point transitions (Tm, see text) were as follows MA: Tm ¼ 55 6.4 C, MI: Tm ¼ 41 2 C, flagelliform: Tm ¼ 44 5.3 C, cylindriform: Tm ¼ 38 2.4 C, pyriform: Tm ¼ 61 3.5 C, acinous: Tm ¼ 62 1.1 C (A, B, C, D modified from Dicko 2004b).

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seven (Fig. 7) silk gland proteins from Nephila edulis at 20% w/v concentration (henceforth % wt.). Each panel shows the spectra at 20 C and 80 C (for experimental details see Dicko et al., 2004b). Three main characteristics emerge from these data. First, silks with different functions appear to have different conformations in solution; second, we observe an inverse temperature transition to a b‐sheet or b‐turn like state; and finally, silk crystallization requires temperatures that are much lower than those required for typical synthetic polymers. The inverse temperature (Urry et al., 1992) is a feature found in all silks; meaning that with increasing temperature rather than unfolding, the structure undergoes a conformational change to another defined structural state (Dicko et al., 2004b,d). For example, in panels (A) and (B) of Fig. 8 both major and minor ampullate silks undergo a conformational change from a helical state toward a b‐sheet‐rich state. On the other hand, panels (D), (E), and (F) show, for the cylindriform, pyriform, and acinous silks, transitions from helical structures (a‐helix for cylindriform and acinous and most likely a 310 helix for pyriform) to b‐sheet/b‐turn structures. The loss of signal intensity suggests that the final spectra at 80 C are in fact building units constitutive of the structure at 20 C. Flagelliform silk in panel (C) further illustrates this phenomenon. Flagelliform silk proteins have been hypothesized to form a b‐spiral, which is a large helix formed of successions of b‐turns; and here we observe for the putative b‐spiral structure at 20 C and at 80 C, the building units that identify that it has b‐turns. A similar transition was observed in the B. mori silk fibroin structure in solution (Canetti et al., 1989; Iizuka and Yang, 1966, 1968; Yang et al., 2004). The midpoint transition temperature observed apparently is rather low (Dicko et al., 2004b), which suggests that the energy required for crystallization is also low, except for the acinous and pyriform silks requiring temperatures over 60 C. This may be linked to the lack of obvious b‐crystalline associated sequences in acinous and pyriform silks. The variation in mid‐point temperature, although not enough to explain the molecular mechanism of the transition, provides a good indicator of the energetic cost to produce semi‐crystalline silk polymers. Helping us in our understanding of the structure–function relationship in silks is our access to silk gene‐sequences. Surprisingly, although we know explicitly the location or absence of b‐structures (Table I), the conformation of the prespun silk solution shows that helical or disordered structures are highly predominant. The putative b‐structures in the preprocessed solution appear to be shielded and constrained into ‘‘nonaggregating’’ structures (Monti et al., 2001; Taddei and Monti, 2005; Yao et al., 2004). In amyloids, and other fiber‐forming proteins, the sequences do not explicitly locate b‐sheet‐forming sequences (Halverson et al., 1991), but

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there is still a high probability (when the right conditions are fulfilled) of undergoing conformational change and aggregation (Dobson, 1999; Jaenicke, 1995). This raises the question of intermediate states and silk aggregation as a folding or misfolding event. The hypothesis that specific intermolecular interactions between hydrophobic surfaces of structural subunits in partially folded intermediates are responsible for the formation of inclusion bodies and protein aggregates is supported by a substantial body of data (Fink, 1998; Jaenicke and Seckler, 1997; Kelly, 1996; Speed et al., 1996). Hydrophobic interactions, resulting from the reluctance of nonpolar groups to be exposed to water, are also considered to be the major driving force for protein folding (Dill, 1990; Fink, 1995). Because proteins often exhibit marginal stabilities (typical Gstab values are 50 kJ/mol; Jaenicke, 1991, 1995), both folding and misfolding are two aspects of the delicate balance of exposed and buried hydrophobic surface, thus illustrating the evolutionary selection for protein flexibility (an important parameter for function, regulation, and degradation) rather than rigidity (one for stability). In other words, the rate of formation and stability of the intermediates on the folding and misfolding pathways, respectively on‐ and off‐pathways, will determine the outcome of the balance of forces acting on the protein. In the case of silk proteins, the misfolding hypothesis (Dobson, 1999; Taubes, 1996) or the presence of natively unfolded structures (Kenney et al., 2002) are both open to discussion. A few points can help to clarify the issue. One example is that the very low chemical enthalpy of silk proteins suggests that the native state is actually the b‐sheet structure and that the observed structures in solution are intermediate or ‘‘denatured’’ states. Another example is the observation that both the competition for extended stabilizing structures (Valluzzi and Jin, 2004) and the local segment conformational polymorphism (Dicko et al., 2005; Wilson et al., 2000) depend predominantly on the local environment of the proteins along the processing pathway (Dicko et al., 2004d). In any case, through the propensity and specificity for aggregation (e.g., by b‐sheet formation in amyloids), proteins, silk, or otherwise, they will precipitate and form disordered aggregates or fibrils under the appropriate conditions (whether in vivo or in vitro; Dobson, 1992; Fandrich et al., 2001; Uversky and Fink, 2004).

C.

Factors Governing the Transition to b‐Structures

Although spider and insect spinning silks are different and their silk types and functions diverse (Craig, 2003; Foelix, 1996), it is strongly

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suggested by histochemical, histological, and ultrastructural evidence that most of these silks are processed in a rather similar way (Akai, 1998; Casem et al., 2002; Kovoor, 1977, 1986, 1987, 1990; Kovoor and Lozez, 1988; Peters and Kovoor, 1991; Tillinghast and Townley, 1994; Vollrath and Knight, 2001). This suggests that a series of common characteristics and parameters are critical for the spinning process. From a gross morphological perspective, regionalization is immediately apparent in the silk‐producing glands (Akai, 1998; Kovoor, 1972; Kovoor and Zylberberg, 1972; Peters, 1955). For example, although the causes of silk conversion in the glands are not yet fully understood, the silk‐spinning solution appears to be subjected to water resorption (Tillinghast et al., 1984; Vollrath et al., 1998), a decrease in pH from about 7 to 4–5 (Dicko et al., 2004d; Knight and Vollrath, 2001; Kovoor, 1987; Magoshi et al., 1994; Vollrath et al., 1998) as well as changes in element composition and concentration in both spiders (Knight and Vollrath, 2001) and silk worms (Liu et al., 1997; Magoshi et al., 1994). The origin of the pH gradient (Dicko et al., 2004d) is still unclear, but has been linked to the presence of a proton pump (Azuma and Ohta, 1998; Vollrath et al., 1998) or acidic and alkaline secretion (Kovoor, 1977, 1987, 1990; Peakall, 1969). The high activity of phosphatases (Tillinghast and Townley, 1994) in epithelial cells suggests that pH may be influenced by the secretion of phosphate ions into the lumen of the gland. Despite limited knowledge on the origin of protons (Hþ) and other alkali (e.g., Kþ, Naþ, Ca2þ) and metal ions (e.g., Cu2þ, Znþ), the location and change in these elements is known (Knight and Vollrath, 2001; Liu et al., 1997; Zhou et al., 2003) and has been correlated to silk protein structures and shapes in vivo (Dicko et al., 2004d; Foo et al., 2006; Ochi et al., 2002a; Terry et al., 2004). Their selective effects have also been tested in vitro (Chen et al., 2002; Dicko et al., 2004a). Overall changes in pH and alkali or metal ions will induce a conformational change to a b‐sheet‐rich state correlated to an increase in solution viscosity (Foo et al., 2006; Kim et al., 2004). The chemical complexity involved in the formation of b‐sheet structures contrasts dramatically with the tight physiological controls used to keep globular proteins folded and protected from a conformational accident. Until recently, it was assumed that in only a ‘‘handful’’ of proteins aggregation was linked to a conformational change to b‐structures (Dobson, 1999). However, careful examination of well‐understood systems showed that amyloid conversion is a common phenomenon (Dobson, 1999, 2001). Why this may have arisen is yet to be discovered, although in a recent survey, Broome and Hecht (2000) found that in proteins a binary arrangement (polar–apolar alternation) is significantly rare compared to

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nonbinary arrangements because the former will inherently produce amyloids. In this respect, silks have evolved one of Nature’s most‐maligned pathogenic molecular mechanisms as a keystone of survival. To control such ‘‘structurally reactive’’ proteins, progressive and judicious modifications are necessary. For example, calcium ions were found to stabilize silk proteins at high concentrations, but at low concentrations to destabilize and promote b‐sheet structures (Magoshi et al., 1994; Zhou et al., 2004). Another judicious effect of ions is the progressive exchange of the stabilizing sodium ions for b‐sheet promoting potassium ions during fiber formation in spiders (Chen et al., 2002; Dicko et al., 2004a; Knight and Vollrath, 2001). Interestingly, similar conditions will produce, in globular or fiber‐forming proteins, the formation of amyloids (Chiti et al., 1999; Dobson, 2001; Fandrich et al., 2001; Prusiner, 1998; Sunde and Blake, 1998). However, for these proteins such conditions would be denaturing and thus would promote aggregation‐induced misfolding. It is unclear whether the aggregation of silk proteins involves unfolded intermediates. However, considering the large diversity of spidroins and fibroins, we may argue that the primitive acinous silks are closer to the modern paradigm of amyloid‐forming mechanisms, and that more derived silks such as major ampullate dragline silks or B. mori fibroin are actively stabilized ‘‘intermediates’’ (or natively unfolded; Kenney et al., 2002) ready to convert to a b‐sheet structure and form amyloids. Silks, in other words, could provide a snapshot of the evolution and stabilization of intermediate states (Dicko C., Kenney J., Bond J., unpublished work). In summary, the physiological control of silk protein conversion shows an ingenious balance of activating and inhibiting mechanisms that are dependent on composition and sequence arrangement (Krejchi et al., 1994). Denaturing effects observed in silks appear to be identical to those found in amyloid‐forming proteins, and they principally alter the competitive outcome of the hydration of nonpolar and polar residues (Anfinsen, 1973; Dill, 1990; Dobson and Karplus, 1999; Kauzmann, 1959). The key differences to amyloids may lie in the hierarchical level of the structures (Muthukumar et al., 1997) involved in the assembly of silks compared to amyloids.

D. Irregular Sequence Units and Heterogeneous Structures Crystallization is the basis of silk strength. But if crystallization were excessive, then the silk would become very brittle rather than being the flexible fiber that it is. The study of model peptides inspired from silk sequences has shown catastrophic and uncontrollable aggregation suggesting

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that control points were missing (Valluzzi et al., 1999, 2002). Detailed scrutiny of the gene sequences (Table I) shows a remarkable phenomenon, namely that a large majority of residues are interchangeable (Hayashi and Lewis, 1998; Sehnal and Zurovec, 2004). This could be argued in two ways: (i) that repeats dominated by residues with small side chains are always functional regardless of substitution or (ii) that the repeats observed with atypical (non alanine, glycine, or serine) residues are in fact true repeats needed to break the symmetry and provide sufficient irregularity to the silk proteins to allow for flexibility, solubility, and the prevention of catastrophic aggregation. In general, heterogeneities in structural materials are often the source of mechanical failure, but specific types also provide ways to disperse energy without failure. For example, some silks, at a microscopic and macroscopic level, are able to form structures such as spherulite inclusions that will develop into elongated cavities in the solid fibers (Akai, 1998; Frische et al., 1998; Robson, 1999; Tanaka et al., 2001). Interestingly, Isobe et al. (2000), in a significant but largely overlooked paper, showed that synthetic Ab1–40 produced spherulites that had the essential features of Alzheimer’s amyloid senile plaques (Kaminsky et al., 2006). In silks, neither the origin nor the function of these macroscopic structures is known, but most likely each is dependent on the conformation and hydropathy of folded silk units (Jin and Kaplan, 2003; Knight and Vollrath, 2002) and apparently serve to strengthen the fiber (Shao et al., 1999) in a way analogous to a ‘‘filled rubber.’’ At the molecular level, the role and effect of the irregular GAAS units on the secondary structure of B. mori silk (Asakura et al., 2002a; Ha et al., 2005) is starting to emerge. These units (or motifs) account for most of the distorted structures observed (Asakura and Yao, 2002; Asakura et al., 2002b) with one important consequence: distorted structures are less prone to aggregation (Valluzzi and Jin, 2004). Other irregularities, such as the unequal sizes of repeats or spacers (Sehnal and Zurovec, 2004), and the presence of tyrosine insertions (Asakura et al., 2005), can lead to increased ‘‘randomness’’ and can promote intermolecular interactions. Overall, irregularities in sequence introduce loops and hairpins that facilitate refolding and packing of the large silk proteins (Asakura et al., 2002a; Ha et al., 2005; Kameda et al., 1999). In less derived glands, such as the acinous glands and some cylindriform glands (Table I), the silk sequences do not show the well‐defined poly‐ (alanine) or poly‐(alanine‐glycine) sequences found in major ampullate silks. But instead, they display a more homogeneously composed gene sequence (Garb and Hayashi, 2005; Hayashi et al., 2004; Tian and Lewis, 2005). Interestingly, silk fibers from the acinous glands seem to exhibit

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mechanical properties comparable to major ampullate dragline silk (Hayashi et al., 2004). The extent to which the animal can permit silk protein sequence heterogeneity is not known, but studies on gene expression and evolution (Craig, 2003; Fedic et al., 2003) suggest that a silk’s amino acid content may dramatically affect liquid crystal flow (Braun and Viney, 2003), silk phase diagrams (Magoshi et al., 1994; Sohn et al., 2004), and molecular mobility (Bonthrone et al., 1992; Kishore et al., 2001; Sapede et al., 2005). New approaches to modeling and peptide design in silks, focusing on their nanostructure and hierarchical chains assembly (Makin et al., 2005; Porter et al., 2005; Vollrath and Porter, 2006; Zhou and Zhang, 2005), hopefully will provide answers to the significant amino acid plasticity of silk proteins and the effect this has on their solution and mechanical behavior. Critically, another important aspect of having heterogeneities is the formation of superstructures such as fibrils. Fibrils or any other structures based on silk crystallization will need heterogeneity to nucleate and grow.

IV. A.

Fibril Assembly: Amyloid Nature of Silk?

Spinning Versus Growth: Length and Time Scale Change

The forces that stabilize amyloid fibrils include specific hydrogen bonding, electrostatic interactions, p–p stacking, and hydrophobic interactions. Importantly, similar types of interactions stabilize the functional native structures of protein molecules (Anfinsen, 1973; Dill, 1990; Dobson and Karplus, 1999; Kauzmann, 1959). In this sense, the conditions that favor native protein folding might also be manipulated to facilitate the formation of amyloid fibrils. It is now well understood that fibril formation requires conformational changes, but the assembly steps may differ from one system to another (Kelly, 1998). For example, aggregation into well‐ordered structures occurs in multiple steps during the formation of b‐lactoglobulin fibrils. First, there is a fast and reversible step followed by an irreversible step involving the formation of nonreversible b‐sheet structures (Arnaudov et al., 2003). Interestingly, the reversible step, which corresponds to a lag in fibril formation, varies from one system to another and most likely depends on the specific kinetic partitioning between the misfolded intermediate and the native state (Dobson, 1999; Jaenicke, 1995; Uversky, 2003). The model of amyloid fibril formation is a nucleation step followed by growth, where the nucleation mechanism dictates the concentration and time dependence of the aggregation (Harper and Lansbury, 1997;

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Lomakin et al., 1996). Surface plasmon resonance studies of the Ab(1–40) b‐amyloid peptide (Cannon et al., 2004) provide even more details of the multiple kinetic steps, and suggest that fibril formation proceeds by reversible addition of a monomer to the tip of the formed fibril, followed by a postbinding, transitional event. A comparable folding mechanism was found in silks. A seminal study by Li et al. (2001) found that in vitro formation of silk fibrils is conformation dependent and occurred via a nucleation mechanism. Although now established as amyloidogenic (Kenney et al., 2002), the nature of the silk fibril assembly remains unclear. Noteworthy is the evidence for a cross‐ nucleation ability of silk proteins, supporting the amyloidogenicity of silk (Lundmark et al., 2005). Inoue et al. (2003) found that silk proteins will form rodlike structures and that those structure will assemble into comblike or fabric‐like superstructure. The scale differences between the rods (nanometers) and the superstructure (micrometers) would suggest that the rod formation is governed by amyloid fibril formation and that the supramolecular arrangement is governed by the properties of the rod (Oroudjev et al., 2002; Putthanarat et al., 2000), namely surface interaction and hydration. Three levels of association could be considered: (i) within the proteins internal b‐strands will organize to form intra b‐sheet structures, (ii) b‐sheets from neighboring molecules will associate to form fibril subunits, and (iii) the fibril subunits will further associate to form larger fibrils or rods. This change in scale and interaction regimes would form the basis to differentiate spinning from fibril growth. This is particularly a propos from a materials point of view, where a multifibrillar composition offers a mechanical advantage over a single fiber of the same cross‐sectional area (Putthanarat et al., 2000). One aspect of the silk fibril formed in solution remains unclear: the apparent absence of the cross‐b structure that characterizes amyloid fibrils.

B. Missing Cross‐b Structure A trademark of amyloid fibrils is their cross‐b structure. This structure is the basis of the repetitive hydrogen‐bonding extension of the fibril (Makin et al., 2005). Cross‐b structures are observed in the silk fibers of some insects (Geddes et al., 1968; Hepburn et al., 1979), although none are observed in spiders or lepidoptera (Craig, 1997). This absence has been explained by the possibility that cross‐b silks or a‐silks may be converted into collinear b‐silks by stretching the fiber and an increased orientation‐ function correlated to the speed at which silk is formed (Riekel et al., 2000).

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A better characterization of fibrils found in various silks could resolve this issue and most likely reveal the conformational criteria involved in the choice of the collinear‐b over the cross‐b structures.

V. Conclusions The spider’s ability to organize, assemble, and partition soluble proteins into semicrystalline fibers relies on firm control during the arrangement of the constituent molecules (or molecular motifs) into tightly folded b‐sheet structures. Hence the animals have been selected to efficiently manage amyloidogenic (i.e., b‐sheet forming) growth under natural, physiological conditions. This distinguishes silks from globular proteins that typically require unusual (marginal) conditions to exhibit b‐sheet driven aggregation (Fandrich et al., 2001). In effect, the evolutionary history of silks with strong selection on these fibers as an integral part of the spider’s reproductive fitness (Vollrath, 2000a) contrasts strongly with the phylogeny of other ‘‘naturally’’ occurring fiber‐forming proteins such as amyloids and prions (Dobson, 1999). While these proteins may also aggregate into fibrils under physiological conditions, their formation is considered the outcome of complex interactions running out of control (Dobson, 2001), thus becoming pathogenic and ultimately lethal for the replication of the protein itself. Combined with the minute quantities of material available, this apparent lack of evolutionary history makes the study of amyloids much more difficult than the study of silk formation where the traits observed reflect molecular optimization rather than conflict and we can examine huge quantities of a highly diverse range of materials that are linked through 400 years of evolution. The intrinsic ability of silk proteins to form b‐sheet structures with stringent checks on premature aggregation is the hallmark of fiber formation. This happens via a progressive protein conformational transition and gel formation mediated by factors such as pH and ionic interactions along the silk production pathway. All research has shown that the environment of the postexpression processing is at least as important as the silk amino acid sequence. After all, it is the spinning process that requires specific superstructures (either micellular and/or rodlike), which themselves are then the foundation of the peculiar (and highly variable) mechanical properties of different fibers. The silk production process must have some built‐in temporal, mechanical, and chemical flexibility. This is evident in the ability of the animal to switch from long‐duration storage of the proteins (in an aqueous solution state) to high‐speed extrusion spinning (into an insoluble fiber with

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properties vastly different from its liquid state). Additionally, the animal can modify the production parameters to create silk fibers with highly diverse mechanical properties. The animal does this by storing the silk in a tightly controlled metastable gel state that is presumably already primed for rapid conversion (perhaps through an intermediate state) to form the fiber. One could compare b‐sheet driven aggregation with a runaway process that Nature does her best to control. Indeed, the ability of nearly all proteins to form a stable aggregate state (albeit mostly through nonphysiological conditions) seems ubiquitous, which suggests that silks may hold the key to understanding amyloidogenicity and its evolution. We conclude that combining the insights coming from studying these two rather different animal products, silks and amyloids, may provide us with answers to questions that are of great interest to both spider and man, albeit for the opposite reasons.

Acknowledgments We thank the Danish SNF (grant 21–00–0485), the British EPSRC (grant GR/NO1538/01) and BBSRC (S12778), the European Commission (grant G5RD‐CT‐2002‐00738), the AFSOR of the United States (grant F49620‐03‐1‐0111), the Institute for Synchrotron Radiation, ISA, Aarhus, Denmark for use of their CD facility through an EC‐Human Potential Program Transnational Access to Major Research Infrastructures (EU Contract No RII3‐CT‐ 2004‐506008). C.D. is supported by St Edmund Hall, Oxford Junior Research Fellowship. J.M.K and C.D thank the State of North Carolina 2003 Biotechnology Instrumentation Initiative for support.

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b‐ROLLS, b‐HELICES, AND OTHER b‐SOLENOID PROTEINS By ANDREY V. KAJAVA* AND ALASDAIR C. STEVEN{ *Centre de Recherches de Biochimie Macromolećulaire, CNRS FRE‐2593, 1919 Route de Mende, 34293 Montpellier Cedex 5, France; { Laboratory of Structural Biology, National Institute of Arthritis, Musculoskeletal, and Skin Diseases, National Institutes of Health, Bethesda, Maryland 20892

I.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. b‐Roll, b‐Helix, and b‐Solenoid: What Are They?. . . . . . . . . . . . . . . . . . . . B. What Are the Distinguishing Structural Features of b‐Solenoids? . . . . C. Stacking of Side Chains: Asparagine Ladders. . . . . . . . . . . . . . . . . . . . . . . . D. Triple‐Stranded b‐Solenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. To Be or Not to Be a b‐Solenoid? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Diversity and Classification of b‐Solenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Nonredundant Set of b‐Solenoid Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . B. Handedness. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Twist . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Relationships Between Handedness and Twist. . . . . . . . . . . . . . . . . . . . . . . E. Cross‐Sectional Shapes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Classification of b‐Solenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Capping and Bulging. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Multistranded b‐Solenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Relationship Between b‐Solenoid Structures and Their Amino Acid Sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. b‐Solenoids Have a Characteristic Amino Acid Composition . . . . . . . . . B. Amino Acid Sequences of b‐Solenoids Have Arrays of Tandem Repeats C. Recurring Conformations and Amino Acid Sequence Motifs of b‐Arcs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Differences Between Sequences of Single‐Stranded and Triple‐Stranded b‐Solenoids. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Examples and Perspectives of Sequence ‐Based Prediction of b‐Solenoid Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Prediction of Amyloidogenic Regions and Structures of Amyloid Fibrils. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Relationship Between b‐Solenoid Structures and Their Functions . . . . . . . . A. Functional Implications of Elongated Shape and Rigidity of b‐Solenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Functional Implications of Highly Regular b‐Solenoid Structures . . . . C. b‐Solenoids as Oligomerization Motifs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. b‐Solenoids as Scaffolds for Multidomain Complexes. . . . . . . . . . . . . . . . VII. Evolution of b‐Solenoid Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Independent Evolutionary Paths of b‐Solenoid Folds . . . . . . . . . . . . . . . . B. Homologous b‐Solenoid Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Perspective. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

ADVANCES IN PROTEIN CHEMISTRY, Vol. 73 DOI: 10.1016/S0065-3233(06)73003-0

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Abstract b‐Rolls and b‐helices belong to a larger group of topologically similar proteins with solenoid folds: because their regular secondary structure elements are exclusively b‐strands, they are referred to as b‐solenoids. The number of b‐solenoids whose structures are known is now large enough to support a systematic analysis. Here we survey the distinguishing structural features of b‐solenoids, also documenting their notable diversity. Appraisal of these structures suggests a classification based on handedness, twist, oligomerization state, and coil shape. In addition, b‐solenoids are distinguished by the number of chains that wind around a common axis: the majority are single‐stranded but there is a recently discovered subset of triple‐stranded b‐solenoids. This survey has revealed some relationships of the amino acid sequences of b‐solenoids with their structures and functions—in particular, the repetitive character of the coil sequences and conformations that recur in tracts of tandem repeats. We have proposed the term b‐arc for the distinctive turns found in b‐solenoids and b‐arch for the corresponding strand‐turn‐strand motifs. The evolutionary mechanisms underlying these proteins are also discussed. This analysis has direct implications for sequence‐based detection, structural prediction, and de novo design of other b‐solenoid proteins. The abundance of virulence factors, toxins and allergens among b‐solenoids, as well as commonalities of b‐solenoids with amyloid fibrils, imply that this class of folds may have a broader role in human diseases than was previously recognized. Thus, identification of genes with putative b‐solenoid domains promises to be a fertile direction in the search for viable targets in the development of new antibiotics and vaccines.

I. Introduction A. b‐Roll, b‐Helix, and b‐Solenoid: What Are They? Over the past few years, a new set of b‐structural proteins has emerged whose folds are based on solenoidal windings of the polypeptide chain. The repeating unit is an individual coil of the solenoid which consists of 12–30 amino acids configured as 2, 3, or 4 b‐strands, together with connecting turns. Each coil in the solenoid has an axial rise of about 4.8 A˚. Corresponding strands in successive coils align to form parallel b‐sheets (Fig. 1). The shortest b‐solenoid on record has three coils (serralysin— Baumann et al., 1993), the longest one has >80 coils [the filamentous hemagglutinin (FHA) of Bordetella pertussis—Kajava et al., 2001], and even

b‐SOLENOID PROTEINS

A

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B

Fig. 1. Schematic representation of a b‐solenoid. The b‐strands are shown as gray arrows. Panel (A) illustrates the hand (in magenta) and twist (in red) of the solenoid. The b‐solenoid shown is right‐handed. Twist is determined as a sense of a virtual helix (red‐dashed line) that connects corresponding reference points in consecutive coils (red points). This b‐solenoid has a left‐handed twist. The larger the twist, the larger the value of a dihedral angle between the vectors (red arrows) connecting the solenoid axis (blue) with the reference points (for details see Section II.B and C). Panel (B) depicts the interior of this b‐solenoid. Apolar (green) and polar (magenta) side chains of interior residues are shown in an atomic space‐filling representation. The front side of the b‐solenoid is omitted to expose the hydrophobic core with greater clarity. The b‐solenoid shown has two ladders of polar residues. The molecular structures in this and other figures were generated with Pymol (DeLano, 2002).

longer ones are likely (Kajava and Steven, 2006). In the interior of these molecules, amino acid side chains are tightly packed, forming a compact, predominantly hydrophobic, core (Fig. 1B). The first structures of this kind were reported in 1993: pectate lyase C from Erwinia chrysanthemi (Yoder et al., 1993) and alkaline protease from Pseudomonas aeruginosa (Baumann et al., 1993). Based on consideration of these crystal structures, the term ‘‘parallel b‐helix’’ was introduced for a fold containing three b‐strands per coil, and ‘‘parallel b‐roll’’ for a fold with two b‐strands per coil (Baumann et al., 1993; Yoder and Jurnak, 1995; Yoder et al., 1993). The epithet ‘‘parallel’’ was intended to emphasize the distinction between these folds and the previously observed helical structure of the antibiotic gramicidin which contains both l‐ and d‐amino acids and

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has an antiparallel b‐sheet (Langs, 1988; Wallace and Ravikumar, 1988). In this classification, b‐strands contain only amino acid residues that have f and c values characteristic of classical b‐structure and in which both the carbonyl and amide groups form interstrand hydrogen bonds with other peptide groups. Subsequently, it was realized that b‐rolls and b‐helices belong to a larger group of topologically similar solenoid proteins whose coils may contain one or several distinct secondary structure elements (a‐helices, b‐strands, 310‐helices, or polyproline II helices; Kobe and Kajava, 2000). b‐Rolls and b‐helices have solely b‐structural repetitive units and, in this classification, both are referred to as b‐solenoids. The close similarity between b‐rolls and b‐helices and potential ambiguity in assigning the starts and ends of b‐strands has led to confusion over whether some newly determined b‐solenoids should be designated as b‐rolls or b‐helices. For example, the C‐terminal domain of glutamate synthase from Synechocystis sp. and the antifreeze protein from Tenebrio molitor were called b‐helices (Liou et al., 2000; van den Heuvel et al., 2003), although they are composed of two‐stranded coils and, therefore, in the original nomenclature (Yoder and Jurnak, 1995), are b‐rolls. Moreover, some recently determined structures have four strands per coil (Dodatko et al., 2004; Hegde et al., 2005); in a strict sense, they are not eligible to be called b‐helices or b‐rolls but they are, nevertheless, b‐solenoids. Here, we will use the term b‐solenoid broadly, as it unites these essentially similar structures in a single structural category, at the same time alluding to their membership of the larger family of solenoid proteins.

B. What Are the Distinguishing Structural Features of b‐Solenoids? Solenoid proteins, with their arrays of repeating motifs, tend to have elongated structures that contrast with the majority of globular proteins whose polypeptide chains follow more complex trajectories (Kobe and Kajava, 2000). In turn, b‐solenoids have several features that distinguish them from other solenoids. First, their coils are composed of all‐b secondary structural elements (Fig. 1A). The coils are generated by the alternation of linear b‐strand segments with tight turns called b‐arcs (Hennetin et al., 2006) or long irregular loops. As a result, the polypeptide chain coils repetitively around the axis of the solenoid. For completeness, we note that not all linear segments are b‐strands. For example, the coils of the C‐terminal domain of bacterial glutamate synthase (Binda et al., 2000; van den Heuvel et al., 2002) have, along with b‐strands, a segment that is overall linear but is made up of several zigzagging b‐arcs. Like b‐strands, it is capable of forming inter‐coil H‐bonds.


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A second distinguishing feature of b‐solenoids is their inter‐coil interactions. Their stacked coils are coordinated by a network of H‐bonds. Usually, the coils in a given b‐solenoid have similar cross‐sectional shapes, an arrangement that allows corresponding strands in successive coils to form parallel b‐sheets. The term b‐arc has been proposed for the turns between strands (Hennetin et al., 2006) and they also stack with inter‐coil H‐bonds, yielding so‐called b‐arcades. The inter‐coil distance (4.8 0.2 A˚) is determined by the distance between H‐bonded b‐strands, which is the same on all sides of the solenoid. Consequently, b‐solenoids are straight and they contrast in this respect with other kinds of solenoids which mostly have curved shapes (Groves and Barford, 1999; Kobe and Kajava, 2000). Another property specific to b‐solenoids is the existence of a well‐ defined boundary between the interior side chains that form the hydrophobic core and the side chains at solvent‐exposed surfaces. This boundary is formed by a tightly packed layer of H‐bonded polypeptide backbones wrapping around the hydrophobic core.

C.

Stacking of Side Chains: Asparagine Ladders

A striking feature of b‐solenoids is the occurrence of stacking interactions among interior side chains (Yoder and Jurnak, 1995; Yoder et al., 1993). Adjacent coils tend to have certain positions occupied by the polar residue— most commonly, asparagine, but also serine, threonine, or histidine (Fig. 1B). Thus, a common motif of parallel b‐helices is the ‘‘asparagine ladder’’ whereby asparagine residues recur at the same interior positions and are arranged so that their side chains stack, forming H‐bonds with each other and with the polypeptide backbone (Fig. 2). Another distinctive stacking interaction involves aromatic side chains configured to give face‐to‐face packing of their aromatic rings (Yoder and Jurnak, 1995).

D. Triple‐Stranded b‐Solenoids All the above‐mentioned proteins have single‐stranded folds based on solenoidal windings of one polypeptide chain. Recently, however, several triple‐stranded b‐helices (alternatively, ‘‘triple‐stranded b‐solenoids’’) have been described in bacteriophage tail proteins (Kanamaru et al., 2002; Smith et al., 2005; Stummeyer et al., 2005; van Raaij et al., 2001). In these structures, three identical chains wind around a common axis and their coils have an axial rise of 14.5 A˚, that is, 3 4.83 A˚ (for details see Sections IV and V.D). In this chapter, triple‐stranded b‐solenoids will be abbreviated as ‘‘TS b‐solenoids,’’ while the term ‘‘b‐solenoid,’’ if not otherwise qualified, will apply to the predominant group of single‐stranded b‐solenoids.

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A

B

C

N

Fig. 2. Ball‐and‐stick representations of two differently oriented asparagine ladders of (A) bl‐arcade taken from the crystal structures of pectate lyase C (Lietzke et al., 1996) and (b) ppl‐arcade taken from UDP‐N‐acetylglucosamine acyltransferase (Raetz and Roderick, 1995). b, l, and so on refer to a one‐letter conformational code (Fig. 10C). The ladders are viewed from within the respective b‐solenoids. The arrow shows the orientation (N‐ to C‐terminal) of the solenoid. Oxygen atoms are in red, nitrogen in blue, and carbon in green. Dotted lines designate H‐bonds of side chains (red) and inter‐coil H‐bonds of the polypeptide backbone (black). Except for the ladder‐forming asparagines, only the backbones of the coils are shown. Panels are reprinted from Hennetin et al. (2006) with the permission of the publisher.

E. To Be or Not to Be a b‐Solenoid? Protein structures are so diverse that it is sometimes difficult to assign them unambiguously to particular structural classes. Such ‘‘borderline’’ cases are, in fact, useful in that they mandate precise definition of the structural classes. In the present context, several proteins have been called ‘‘b‐helical’’ although, in a strict sense, they do not fit the definitions of b‐helices or b‐solenoids. For example, Perutz et al. (2002) proposed a ‘‘water‐ filled nanotube’’ model for amyloid fibrils formed as polymers of the Asp2Gln15Lys2 peptide. This model has been called b‐helical (Kishimoto et al., 2004; Merlino et al., 2006), but it differs from known b‐helices in that: (i) it has circular coils formed by uniform deformation of the peptide b‐conformation with no turns or linear b‐strands, as are usually observed in b‐solenoids; and (ii) it envisages a tubular structure with a water‐filled axial lumen instead of the water‐excluding core with tightly packed side chains that is characteristic of b‐solenoids. Another example is a solenoid domain observed in type I insulin‐like growth factor receptor (IGFR) and epidermal growth factor receptor (EGFR) (Cho and Leahy, 2002; Garrett et al., 1998). In this solenoid, one half of the coil consists of b‐strands that have an axial rise of about 4.8 A˚ and form a parallel b‐sheet. The other half of the coil has an irregular, though extended, conformation. These segments are wider than b‐strands


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and only come as close to each other as 5.5–6.5 A˚—not close enough to allow the formation of inter‐coil backbone H‐bonds. This difference in the inter‐ coil spacing on opposite sides of the solenoid in IGFR or EGFR domains disqualifies them as b‐solenoids, as defined above. The slightly curved structures of the IGFR and EGFR solenoids resemble bacterial leucine‐rich repeat (LRR) proteins (Evdokimov et al., 2001; Kajava, 1998), although their coils do not have a consensus sequence characteristic of LRRs.

II.

Diversity and Classification of b‐Solenoids A. Nonredundant Set of b‐Solenoid Proteins

Since 1993 when the first crystal structures of b‐solenoids were reported, the number of solved structures of this kind has grown steadily. Today, the protein data bank (PDB) (Berman et al., 2000) has over 100 such entries. However, some of these are for proteins with the same or similar amino acid sequences. In surveying them, we found that b‐solenoids with sequence identity above 18% tend to have similar structures, while below this cutoff, the structures may diverge (Hennetin et al., 2006). By this criterion, we compiled a nonredundant set of b‐solenoid structures. It contains 38 representative crystal structures (Tables I and II) and includes several groups of homologous b‐solenoids: (i) bacterial, plant, and fungal enzymes for the degradation and/or modification of polysaccharides (14 proteins); (ii) bacterial transferases (9 proteins); (iii) subunits of bacterial oxidoreductase (2 proteins); and (iv) eukaryotic RP2 and C‐CAP proteins (2 proteins). The majority are bacterial proteins (31 of 38), but there are also 6 from eukaryotes and 1 viral protein. In this section, we review this nonredundant set, document that its members exhibit substantial diversity, and suggest a subclassification based on characteristic features.

B.

Handedness

One basic structural property of a solenoid is its hand (Fig. 1A), that is, the sense with which the chain winds around the molecular axis. Most solenoids, especially the known a‐ and a/b‐solenoids, are right‐ handed (Groves and Barford, 1999; Kobe and Kajava, 2000). b‐Solenoids depart from this trend in that they encompass both right‐ and left‐handed forms (Tables I and II, Fig. 3). Right‐handed structures predominate (70%), but left‐handed structures account for a significant portion of the set (30%). It is noteworthy that the coils of left‐handed b‐solenoids have not more than 18 residues, while the coils of right‐handed b‐solenoids are mostly longer (although some of them are also shorter than 18 residues).

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Table I Nonredundant Set of Known Right‐Handed b‐Solenoid Structures Name

Origin

PDB Resolution code (A˚) References

Bacterial, plant, and fungal enzymes for degradation/modification of polysaccharides: left‐hand‐twisted b‐solenoid with L‐type cross section Pectate lyase C Erwinia 1AIR 2.2 Lietzke chrysanthemi et al. (1996) Pectate lyase A Erwinia 1PE9 1.6 Dehdashti chrysanthemi et al. (2003) Pectate lyase Pel9A Erwinia 1RU4 1.6 Jenkins chrysanthemi et al. (2004) Pel‐15 pectate lyase Bacillus sp. Strain 1EE6 2.3 Akita Ksm‐P15 et al. (2001) Pectin lyase B Aspergillus niger 1QCX 1.7 Vitali et al. (1998) Polygalacturonase Erwinia carotovora 1BHE 1.9 Pickersgill subsp. carotovora et al. (1998) Chondroitinase B Flavobacterium 1DBG 1.7 Huang heparinium et al. (1999) Plant pectin Dancus carota 1GQ8 1.75 Johansson methylesterase et al. (2002) Polygalacturonase I Aspergillus niger 1NHC 1.7 van Pouderoyen et al. (2003) Dex49A dextranase Penicillium 1OGO 1.65 Larsson minioluteum et al. (2003) Rhamnogalacturonase A Aspergillus aculeatus 1RMG 2.0 Petersen et al. (1997) Major pollen allergen Juniperus ashei 1PXZ 1.7 Czerwinski et al. (2005) Jun a1 iota‐Carrageenase Alteromonas fortis 1H80 1.6 Michel et al. (2001) Pectin methylesterase Erwinia chrysanthemi 1Q JV 2.37 Jenkins et al. (2001) Viral enzyme for degradation of polysaccharides: trimer of b‐solenoids with L‐type cross section and a small left‐handed twist Tailspike Salmonella 1QQ1 1.8 Schuler endorhamnosidase typhimurium et al. (2000) phage P22 Bacterial virulence factor: b‐solenoid with L‐, T‐type cross sections and left‐handed twist P.69 pertactin Bordetella pertussis 1DAB 2.5 Emsley et al. (1996) (continued )


TABLE I Name

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(continued)

Origin

PDB Resolution code (A˚) References

Bacterial proteins: b‐solenoids with T‐type cross section and left‐handed twist Filamentous Bordetella pertussis 1RWR 1.72 Clantin hemagglutinin et al. (2004) Hemoglobin protease Escherichia coli 1WXR 2.2 Otto et al. (2005) MinC cell division Thermotoga 1HF2 2.2 Cordell inhibitor maritima et al. (2001) Subunit of bacterial oxidoreductase: b‐solenoids with B‐type cross section and a small left‐handed twist C‐terminal domain of Azospirillum 1EAO 3.0 Binda glutamate synthase brasilense et al. (2000) C‐terminal domain of Synechocystis sp. 1OFD 2.0 Van den Heuvel glutamate synthase et al. (2003) Bacterial proteins: b‐solenoids with O‐type cross section and a small left‐handed twist PrtC protease C Erwinia 1K7I 1.59 Hege and Chrysanthem Baumann (2001) Stabilizer of iron Escherichia coli 1VH4 1.5 Badger transporter SufD et al. (2005) b‐Solenoids with R‐type cross section and a small left‐handed twist Antifreeze protein Tenebrio molitor 1EZG 1.4 Liou et al. (2000) MfpA inhibitor of DNA gyrase Mycobacterium 2BM5 2.0 Hegde tuberculosis et al. (2005) Domain of cyclase‐associated Saccharomyces 1K4Z 2.3 Dodatko protein cerevisiae et al. (2004) Retinitis pigmentosa Homo sapiens 2BX6 2.1 Kuhnel protein 2 (RP2) et al. (2006)

C. Twist Another basic property of solenoids is their twist. This parameter reflects the fact that successive coils are not stacked exactly above one another but with a small angular offset. Twist may be defined in relation to reference points at corresponding positions in consecutive coils (Fig. 1A). When connected, these points form a helix. The sense of the twist is given by the hand of this helix as it winds around the solenoid axis. Note that the definition of protein solenoid twist (Kobe and Kajava, 2000) differs from the usual definition of the twist in b‐sheets which is defined by the twist of b‐strands

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Table II Nonredundant Set of Known Left‐Handed b‐Solenoid Structures Name

Origin

PDB code

Resolution (A˚)

References

Bacterial transferases: flat trimeric b‐solenoids with T‐type cross section N‐Acetyl‐glucosamine Streptococcus 1HM9 1.75 Sulzenbacher 1‐phosphate pneumoniae et al. (2001) uridyltransferase UDP‐N‐acetylglucosamine Helicobacter 1J2Z 2.1 Lee and acyltransferase pylori Suh (2003) Tetrahydrodipicolinate Mycobacterium 1KGQ 2.0 Beaman N‐succinyltransferase bovis et al. (2002) Streptogramin A Enterococcus 1MR7 1.8 Kehoe acetyltransferase faecium et al. (2003) Maltose‐O‐acetyltransferase Escherichia coli 1OCX 2.15 Lo Leggio et al. (2003) Anhydrase Methanosarcina 1QRE 1.46 Iverson thermophila et al. (2000) Serine acetyltransferase Escherichia coli 1T3D 2.2 Pye et al. (2004) Ferripyochelin binding Pyrococcus 1V3W 1.5 Jeyakanthan protein horikoshii and Tahirov (2003) Bacterial transferases: flat b‐solenoid with T‐type cross section ADP‐glucose Solanum 1YP2 2.11 Jin et al. (2005) pyrophosphorylase tuberosum Insect antifreeze protein: flat b‐solenoid with T‐type cross section Antifreeze protein Choristoneura 1L0S 2.3 Leinala fumiferana et al. (2002) Bacterial adhesin: right‐hand‐twisted, trimeric b‐solenoids with O‐type cross section YadA adhesin Yersinia 1P9H 1.6 Nummelin enterocolitica et al. (2004)

when viewed along the polypeptide chain (Chothia, 1973). The vast majority of b‐sheets in known structures are twisted (right‐handed in the conventional definition when viewed along the chain; Salemme and Weatherford, 1981). This trend suggests that such a twist is energetically favorable compared to b‐sheets that have the opposite twist or are flat. b‐Solenoids conform to this rule: the majority of their parallel b‐sheets are right‐hand‐twisted along the chain and left‐hand‐twisted along the solenoid axis. Indeed, 26 of 38 b‐solenoids of the nonredundant set are left‐ hand‐twisted. Of the remainder, 11 are flat and only 1—YadA adhesin (Nummelin et al., 2004)—is an unusual right‐hand‐twisted b‐solenoid (Tables I and II).

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Pectate lyase C

P.69 pertactin

Tailspike endorhamnosidase

MinC cell division inhibitor

PrtC protease C

Glutamate synthase

Fig. 3.

(continued )

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Stabilizer of iron transporter SufD

Antifreeze protein

Cyclase-associated protein

MfpA inhibitor of DNA gyrase

Antifreeze protein

YadA adhesin

N-acetyl-glucosamine 1-phosphate uridyltransferase

Fig. 3. A gallery of structures representative of the distinct b‐solenoid groups shown in Tables I and II. The b‐solenoid domains are in blue and other domains are in yellow. In the oligomeric structures, only one subunit is colored while the other ones are gray. Small ligand molecules are shown in the ball‐and‐stick representation and colored magenta.


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D. Relationships Between Handedness and Twist It has been suggested that the hand of b‐solenoids may depend on both the twist of the b‐strands and side chain packing in the vicinity of the b‐arc turns (Kobe and Kajava, 2000). Analysis of known b‐solenoids (Tables I and II) supports this suggestion. All left‐hand‐twisted b‐solenoids are right‐handed, while the only one with a right‐hand‐twist—YadA adhesin (Nummelin et al., 2004)—is left‐handed. Flat b‐solenoids may be either right‐ or left‐handed. Depending on their hand, b‐solenoids also have preferential b‐arch conformations (Section V.C). Interestingly, most of the left‐handed b‐solenoids (bacterial transferases, YadA adhesin) form trimeric structures (Fig. 3). At the same time, all right‐handed b‐solenoids except phage P22 tailspike (Schuler et al., 2000) are monomers. This correlation suggests that folding of b‐solenoids in an a priori less favorable left‐handed arrangement may be facilitated by inter‐solenoid interactions. Otherwise, relationships between amino acid sequences and b‐solenoid handedness and twist are not yet well established.

E. Cross‐Sectional Shapes The structural diversity of b‐solenoids mainly reflects variations in the shape of the coils as viewed along the solenoid axis. While coil shape is, to a good approximation, maintained consistently within a given b‐solenoid, coils from different b‐solenoids vary markedly. Coils differ in their lengths (12–30 amino acids), in the number of b‐strands per coil, and in the type and order of b‐arc conformations. These variations generate molecules whose cross‐sectional shapes resemble the letter ‘‘L’’ (L‐type), the letter ‘‘B’’ (B‐type), an oval (O‐type), a rectangle (R‐type), or a triangle (T‐type) (Fig. 4). In the nonredundant set, L‐type b‐solenoids are the most prevalent and include various bacterial, plant, and fungal enzymes for the degradation and modification of polysaccharides (Akita et al., 2001; Czerwinski et al., 2005; Dehdashti et al., 2003; Jenkins et al., 2001, 2004; Johansson et al., 2002; Larsson et al., 2003; Michel et al., 2001; Petersen et al., 1997; Pickersgill et al., 1998; van Pouderoyen et al., 2003; Vitali et al., 1998; Yoder et al., 1993), phage P22 tailspike endorhamnosidase (Schuler et al., 2000), and the P.69 pertactin virulence factor from B. pertussis (Emsley et al., 1996). T‐type b‐solenoids constitute the other well‐populated category that includes several trimeric bacterial transferases (Beaman et al., 2002; Iverson et al., 2000; Kehoe et al., 2003; Kisker et al., 1996; Lee and Suh, 2003; Lo Leggio et al., 2003; Sulzenbacher et al., 2001), antifreeze protein from Choristoneura fumiferana (Leinala et al., 2002), and several bacterial virulence factors (Clantin et al., 2004; Emsley et al., 1996; Otto et al., 2005).

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O-type

R-type

T-type

B-type

L-type

Fig. 4. Representative structures of the distinct types of coil shapes, viewed along the b‐solenoid axis. The structures shown are limited to two coils. The b‐strands are shown as red arrows connected by loops. The side chains are shown in the ball‐and‐stick representation and colored gray.

Four proteins are R‐type b‐solenoids: antifreeze protein from T. molitor (Liou et al., 2000), DNA gyrase inhibitor MfpA from Mycobacterium tuberculosis (Hegde et al., 2005), and two homologous eukaryotic proteins, RP2 and C‐CAP (Dodatko et al., 2004; Kuhnel et al., 2006). They are quite similar in shape to O‐type b‐solenoids [b‐rolls of PrtC protease from E. chrysanthemi (Hege and Baumann, 2001) and of iron transporter stabilizer SufD from Escherichia coli (Badger et al., 2005)]. The main difference between them is that O‐type coils have two b‐strands packed against each other and linked by two short b‐arcs with fewer than four residues on the exterior surface, while R‐type coils have longer and linear regions (b‐strands) instead of these b‐arcs (Fig. 4). Finally, C‐terminal domains of bacterial glutamate synthase (Binda et al., 2000; van den Heuvel et al., 2002) have b‐solenoids with an unusual B‐type coil. In this structure, one side of the solenoid is made up of several zigzagging b‐arcs that produce an approximately linear segment capable of forming inter‐coil H‐bonds. As mentioned above, coil shape is usually maintained within a given b‐solenoid. However, some coils in a given solenoid may have long loops instead of tight b‐arcs but otherwise retain the overall shape of the coil. Occasionally, different coil shapes are found in the same molecule. For example, P.69 pertactin has several L‐type coils followed by T‐type and O‐type coils (Emsley et al., 1996).

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F. Classification of b‐Solenoids Appraisal of known single‐stranded b‐solenoids suggests a classification based on the following characteristics. The first ones are handedness and twist. So far, four types of b‐solenoids have been observed: right‐handed with a left‐hand‐twist; left‐handed with a right‐hand‐twist; left‐handed, flat (untwisted); and right‐handed, flat (Tables I and II). The next distinguishing feature is coil shape (Fig. 4), for which we recognize five types: B‐, L‐, O‐, R‐, and T‐type (Tables I and II). Among known structures, L‐, B‐, and R‐type solenoids are exclusively right‐handed. Moreover, all of them, except for the flat R‐type b‐solenoid of antifreeze protein from T. molitor (Liou et al., 2000), are left‐hand‐twisted. T‐ and O‐type b‐solenoids may be either right‐ or left‐handed. Finally, we consider oligomerization state (Tables I and II). Most b‐solenoids are monomers, but there are some trimers with right‐handed L‐type (Schuler et al., 2000) and left‐handed T‐type solenoids (Beaman et al., 2002; Kehoe et al., 2003; Kisker et al., 1996; Lo Leggio et al., 2003; Sulzenbacher et al., 2001). The trimers are formed by lateral, in‐register, association of the individual solenoids (Figs. 3 and 5). There is also one

Bacterial transferase

Tailspike endorhamnosidase

YadA adhesin

Fig. 5. Axial projections of b‐solenoid trimers formed by lateral, in‐register, association of the individual solenoids. Only one coil of each solenoid is shown. Different subunits of a trimer are colored in different colors. Ligands of the bacterial transferase are shown in green.

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example of lateral interactions between short T‐type b‐solenoid domains in the dimer of bacterial cell division inhibitor MinC (Cordell et al., 2001). In other dimeric structures, b‐solenoids associate either via tail‐to‐tail interactions of ‘‘open’’ terminal coils as observed with the O‐type solenoid of iron transporter stabilizer, SufD (Badger et al., 2005); by interactions mediated by the capping domains of b‐solenoids as in MfpA inhibitor of DNA gyrase (Hegde et al., 2005); or by swapping of capping domains as in cyclase‐associated protein (CAP) (Dodatko et al., 2004) (Fig. 3).

III.

Capping and Bulging

If a solenoid ends abruptly, the terminal coils with their mutually complementary surfaces and hydrophobic cores are exposed to the solvent. This situation would appear to have a high potential for head‐to‐tail interactions that might lead to polymerization into fibrils of indefinite length. However, it turns out that most protein solenoids are N‐ and/or C‐terminally capped—possibly, to forestall such polymerization. The capping structures come in many forms. The simplest cap is a coil that resembles the other coils of its solenoid but is, however, rich in polar and charged residues that shield the hydrophobic core. Such capping is exemplified in the N‐terminal coil of P.69 pertactin (Emsley et al., 1996). To interrupt the H‐bonding complementarity of the coil backbone, the terminal coil may also contain proline residues and/or b‐bulges (Richardson and Richardson, 2002), as in the antifreeze solenoids (Leinala et al., 2002; Liou et al., 2000). Frequently, the capping coils are rich in bulky aromatic residues such as Tyr or Trp whose side chains overfill the coil interior, preventing other coils from forming H‐bonds with the terminal one. The aromatic residues may also provide an additional stabilization for the capping structure. The second frequently occurring cap motif is an amphipathic segment of polypeptide chain that crosses over the hydrophobic core of the terminal coil. The apolar side of this segment faces the interior while the polar side is exposed to solvent. This segment may have an extended b‐structure conformation as in the N‐terminal coil of bacterial cell division inhibitor MinC (Cordell et al., 2001) and the C‐terminal cap of hemoglobin protease (Otto et al., 2005), or an a‐helix as in pectate lyases (Lietzke et al., 1996). A third kind of capping motif consists of a hydrophilic b‐strand that pairs with a b‐strand of the terminal coil to form an antiparallel b‐hairpin (Cordell et al., 2001; Dodatko et al., 2004; Otto et al., 2005). Finally, caps may consist of globular domains (Clantin et al., 2004; Emsley et al., 1996; Hege and Baumann, 2001). Another possible role of globular caps could be to initiate correct folding of the solenoid (Clantin et al., 2004; Oliver et al., 2003).

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A

B 1048

1

Fig. 6. Terminal capping and lateral bulging of globular domains in the b‐solenoid of the hemoglobin protease from E. coli (Otto et al., 2005). The b‐solenoid domains are shown in blue and the remaining regions in dark yellow. (A) Ribbon diagram of the 3D structure and (B) linear map of the domain distribution within the amino acid sequence.

Bulging of globular domains from the b‐solenoid shaft also bears on the potential of abruptly terminated solenoids to engage in head‐to‐tail interactions. Two b‐solenoid domains may be separated by hundreds of residues in an amino acid sequence but combine in the 3D structure to form a single fused b‐solenoid (Fig. 6). The intervening sequences are either folded into globular domains or form long irregular loops that bulge out from the wall of the solenoid (Otto et al., 2005). The head‐to‐tail interaction between coils is strong enough to preserve the integrity of the solenoid structure, even after proteolytic cleavage of loops in turn sites (Makhov et al., 1994). Bioinformatic analysis of the amino acid sequences of bacterial virulence factors reveals a number of such b‐solenoid domains separated along the chain by lengthy regions (Kajava and Steven, 2006). This observation suggests that b‐solenoids with adjoining globular domains may be quite common.

IV.

Multistranded b‐Solenoids

Most known b‐solenoids are monomers. Recently, however, several triple‐stranded (TS) b‐solenoids have been described in which the three chains wind around a common axis. The three monomers are related by

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threefold rotational symmetry and their coils have an axial rise of 14.5 A˚ (3 4.83 A˚) (Fig. 7). All these solenoids are right‐handed with a slight left‐handed twist. To date, this unusual fold has been found only in bacteriophage tail proteins. It was initially proposed for part of the T4 short tail‐fiber gp12, based on EM data and analysis of sequence repeats (Makhov et al., 1993). Subsequently, van Raaij et al. (2001) identified a crystallizable fragment of gp12 and determined its structure to 1.9‐A˚ resolution, finding three motifs including a well‐ordered segment of TS b‐helix. TS b‐solenoids have since been found in the T4 cell‐puncturing device gp5 (Kanamaru et al., 2002), E. coli K1 bacteriophage K1F endosialidase (Stummeyer et al., 2005), and group A streptococcal phage‐encoded hyaluronidase HylP1 (Smith et al., 2005). A very short motif of this kind with only one coil from three b‐strands is found in the tailspike protein gp9 of phage P22 (Steinbacher et al., 1994). This motif has also been proposed to occur in the T4 long tail‐fibers (Cerritelli et al., 1996) and it may well be present in many other phage tail‐fibers.

A

B

T4 short tail-fiber gp12

gp12

T4 cell-puncturing device gp5

Hyaluronidase HylP1

Fig. 7. Triple‐stranded (TS) b‐solenoids. (A) Ribbon diagram of a fragment of the T4 short tail‐fiber gp12 (van Raaij et al., 2001) and (B) cross‐sectional shapes of the TS b‐solenoids. Repetitive turn‐strand elements of each individual chain of the TS b‐solenoids are rendered in different colors.


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On account of their threefold axial symmetry, the cross‐sectional shapes of these structures are, essentially, equilateral triangles (Fig. 7). Three kinds of triangles have been observed. First, T4 short tail‐fiber gp12 (van Raaij et al., 2001), hyaluronidase HylP1 (Smith et al., 2005), and K1F endosialidase (Stummeyer et al., 2005) have small equilateral triangles formed by sides of six residues. Each side has a three‐residue b‐strand with two of the side chains located inside the structure and a three‐residue b‐arc. Second, larger equilateral triangles with sides of eight residues were found in T4 cell‐puncturing device gp5 (Kanamaru et al., 2002) and also in K1F endosialidase (Stummeyer et al., 2005). The sides of these triangles consist of five‐residue b‐strands (with three interior residues) connected by three‐residue b‐arcs. The third type of cross section, in hyaluronidase HylP1 (Smith et al., 2005), represents a ‘‘collapsed’’ triangle with 11‐ to 20‐residue sides. Each side contains a 6‐residue constant part of a b‐strand with an inverted b‐arc, similar to one previously observed in a single‐ stranded L‐type b‐solenoid (Hennetin et al., 2006; Jenkins and Pickersgill, 2001; Yoder and Jurnak, 1995), and a variable part representing a 5‐ to 14‐residue loop (Fig. 7). Another characteristic of TS b‐solenoids is their degree of structural regularity. Those of T4 gp12 and gp5 are extremely regular (Kanamaru et al., 2002; van Raaij et al., 2001), being generated by repetition of near‐ identical coils of 18 and 24 residues, respectively. In contrast, the structures of K1F endosialidase (Stummeyer et al., 2005) and hyaluronidase HylP1 (Smith et al., 2005) contain coils of varying lengths that generate solenoids of correspondingly variable width. Usually, TS b‐solenoids represent only parts of larger multidomain proteins. Other trimeric motifs found in these proteins include a‐helical coiled coils, TS b‐spirals, trimeric bundles of single‐stranded b‐solenoids, and irregular globular structures. Some of these domains may be needed for correct folding of the TS b‐solenoid. A different kind of multistranded b‐solenoid is represented by the antiparallel double‐stranded b‐helix of cupin proteins (Cleasby et al., 1996; Dunwell et al., 2004). Cupins form a diverse family of enzymes that are typically dimers with a dyad axis. Although the structure looks irregular, on close inspection one can identify a continuous b‐sandwich spanning the dimer (Fig. 8). In each monomer, there are about three coils of double‐stranded b‐helix (b‐solenoid) with one central coil flanked by two coils heading in the opposite direction. As a result, the b‐strands of the coils form antiparallel b‐sheets. The dimers are united via inter‐coil H‐bonding of the N‐termini while the C‐terminal ends are slightly open, so that the b‐structure resembles a funnel. Unlike other kinds of b‐solenoids that, in principle, can be very long, this fold appears to be limited in its axial dimension.

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A

B

Fig. 8. Ribbon diagrams showing (A) lateral and (B) axial views of the antiparallel double‐stranded b‐solenoid from bacterial protein YhcH (Teplyakov et al., 2005). The b‐strands of this dimeric b‐solenoid are colored in blue and magenta.

V.

Relationship Between b‐Solenoid Structures and Their Amino Acid Sequences A. b‐Solenoids Have a Characteristic Amino Acid Composition

Analysis of the amino acid compositions of b‐solenoid domains shows them to be enriched in glycine, valine, asparagine, threonine, and aspartic acid while the incidence of leucine, glutamic acid, lysine, arginine, and proline is lower than average (Fig. 9). The amino acid compositions of single‐stranded and TS b‐solenoids are similar. Some of these variations appear explicable: (i) the preference for valine and threonine correlates with the high b‐sheet propensities of Cb‐branched side chains (Chou and Fasman, 1974), (ii) the high incidence of asparagine can be explained by the presence of the asparagine ladders in b‐solenoids (Yoder and Jurnak, 1995), (iii) glycine is frequently required in b‐arcs (Hennetin et al., 2006), and (iv) proline is poorly represented because this residue destabilizes b‐sheets. Overall, there is a higher incidence of acidic residues (mainly asparagine) than of basic residues in b‐solenoids. Moreover, it is rare to observe charged residues of both signs in these proteins. As yet, explanations for these empirical observations are not apparent.

B.

Amino Acid Sequences of b‐Solenoids Have Arrays of Tandem Repeats

In view of the repetitive 3D structure of b‐solenoids, one might expect their amino acid sequences also to be repetitive. Indeed, some b‐solenoids


75

Fig. 9. Histogram of amino acid composition of b‐solenoid domains from the nonredundant set of known crystal structures (hatched bars) compared to the average one (filled bars) computed from Swiss‐Prot database (Gasteiger et al., 2003). The amino acids are denoted by one‐letter code and are grouped by the following categories (left‐to‐right): amino acid residues whose incidence is respectively unchanged, increased, and decreased in b‐solenoids.

do contain evident sequence repeats. The length of sequence repeat usually matches the length of the coil. In some cases, when the coil cross section has two‐, three‐, or fourfold symmetry, the repeats may correspond to the basic elements of this symmetry and be two, three, or four times shorter than the coil. Among known b‐solenoids, the antifreeze protein from T. molitor (Liou et al., 2000) has the shortest coil of 12 residues formed by 4 similar structural elements. As a result, this protein has a tandem array of 3‐residue repeats with a consensus sequence (A,S,C)‐x‐x. The longest repeats are found, for example, in pectate lyases (Yoder and Jurnak, 1995) or the iron transporter stabilizer SufD (Badger et al., 2005) which have 25–30 residues. The number of repeats in one protein varies from 3 in serralysin (Hege and Baumann, 2001) to about 100 in some autotransporter proteins (Kajava and Steven, 2006). b‐Solenoid repeats usually have several x or xx sequence patterns that correspond to the b‐strands (here, denotes an apolar residue, and x is mostly polar but can be any residue except proline). The middle ‐position in xx usually has a bulky apolar residue, while ‐residues in positions close to turns are often alanine, glycine, serine, or threonine. These positions are also occupied by asparagine residues that stack to form H‐bonded ladders inside the b‐solenoid. The strand‐associated x and xx patterns are interrupted by regions enriched in polar residues and glycine (Hennetin et al., 2006). These are regions of turns and loops. The long loops frequently contain proline residues. In several b‐solenoids, the alternation of apolar and polar residues that is typical for b‐strands is not well observed and ‘‘outside’’ positions are occupied by apolar residues.

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These proteins form dimers or trimers and the ‘‘outside’’ b‐strand residues of the monomer are involved in stabilizing the oligomer. The repeat sequences in solenoid proteins are often so imperfect that it is difficult to identify them without knowledge of the 3D structure or other supplementary information. In highly regular b‐solenoids, such as antifreeze proteins (Leinala et al., 2002; Liou et al., 2000) or YadA adhesin (Nummelin et al., 2004), the repeats are of the same length and sequence similarity between the repeats is discernible (so‐called overt repeats). Frequently, however, the repeats in a given b‐solenoid vary in length because additional residues may be inserted at turn sites without disrupting the pattern of b‐strands that stabilizes the structure (Steinbacher et al., 1994; Yoder et al., 1993). In these cases, the structural repeats are not apparent as sequence repeats (covert repeats). Previous studies (Heffron et al., 1998; Kajava, 1998; Kajava and Steven, 2006) have shown that it is possible to detect covert repeats by applying a sequence profile search (Bucher et al., 1996; Gribskov et al., 1987). The profile method provides a sensitive tool for detecting distantly related members of protein families (Gribskov et al., 1987). A profile combines the information content of a family of sequences. For each position, it contains scoring information for every residue in the alignment, as well as on gap creation and gap extension. Usually, the initial repeat profile is constructed based on the alignment of a few overt repeats and is then edited, taking into consideration information from general knowledge of b‐solenoid architecture or feedback from molecular modeling (Kajava and Steven, 2006). It has also been established that a sequence profile spanning three repeats affords a more sensitive probe of noisy data than a single repeat; moreover, a single repeat of this size would be unlikely to form a stable structure on its own (Kajava, 1998). Another computational approach for detecting b‐solenoid sequences is implemented in a program called BETAWRAP (Bradley et al., 2001). This approach aims to identify b‐solenoid sequences by using hydrophobic‐ residue sequence patterns of strand‐turn‐strand regions that were learned from non‐b‐solenoid structures. This method also takes into consideration the repetitive character of these patterns in b‐solenoids. Unlike the sequence profile approaches, BETAWRAP can make ab initio predictions of b‐solenoid domains. However, it is less sensitive than the profile search and, sometimes, cannot distinguish b‐solenoids from other solenoids (A. V. K., unpublished observation) such as, for example, LRR proteins (Kobe and Deisenhofer, 1994; Kobe and Kajava, 2001). The latest modification of BETAWRAP algorithm, which is called BETAWRAPPRO (McDonnell et al., 2006), employs additional data provided by sequence profiles and this improves the results of b‐solenoid predictions.


C.

77

Recurring Conformations and Amino Acid Sequence Motifs of b‐Arcs

Strand‐turn‐strand motifs in b‐solenoids differ fundamentally from those found in globular proteins. In globular structures, two adjacent strands with an intervening b‐turn form an antiparallel structure called a b‐hairpin (Fig. 10A). In b‐solenoids, the polypeptide chain also folds back on itself, but the flanking b‐strands make contact via their side chains rather than interacting via H‐bonds of the backbone (Fig. 10A). As a result, consecutive strands find themselves in two different, parallel, b‐sheets. The latter strand‐turn‐strand structure is called a b‐arch, and its turn, a b‐arc (Hennetin et al., 2006; Yoder and Jurnak, 1995). In b‐solenoids, b‐arches stack in‐register to form b‐arcades which have two parallel b‐sheets assembled from corresponding strands in successive layers. The geometry of a b‐strand is such that, in a b‐solenoid, its interior and exterior side chains alternate along the chain (Fig. 10B; an interior residue has its side chain buried inside the structure). A discontinuity in this pattern indicates the presence of a b‐arc. The b‐arc can be defined as a region between two b‐strands, each representing a continuous run of three or more residues in b‐conformation that starts and ends with interior residues (Fig. 10B; Hennetin et al., 2006). Despite the apparent irregularity of b‐arcs, they have a limited number of standard conformations that recur in different coils of the same molecule and in proteins that are quite different from both functional and phylogenetic points of view. One of the most frequently occurring arcs consists of two residues in the bl conformation (Hennetin et al., 2006; Jenkins and Pickersgill, 2001; Yoder and Jurnak, 1995; Fig. 11). Here, the letters b, l, and also a, d, g, e, p indicate backbone conformations in corresponding regions of the Ramachandran map (Fig. 10C). The next most abundant arc has three residues in the ppl conformation (Hennetin et al., 2006). Four‐residue arcs do not have such a strong preference for one particular conformation, but several recurring conformations have been identified. Among them are the bepl/bebl, gbpl, and gbeb conformations (Fig. 11). All side chains of the small two‐ or three‐residue arcs are exposed to solvent, while large arcs of more than four residues have one residue in the middle that is buried inside the arch (Fig. 12). From this point of view, four‐residue arcs represent a transition point: gbpl‐like arcs have one interior residue and the other arcs have at least one conserved glycine residue that is partially buried. The most frequently occurring five‐residue arcs have the blbbl conformation. These arcs have one residue in the middle in b conformation that is buried (Figs. 11 and 12). The carbonyl and amide groups of this residue form interstrand H‐bonds with the peptide groups of the corresponding residues in adjacent coils and it can be considered as a

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A

B

2B2 A3 2B3

2B1 1B3

A2

1B1

A1 1B2 C +180 b

p

d I 0

g a

Y

e −180 −180

0 F

+180

Fig. 10. (A) Schematic diagrams of stacks of two b‐hairpins (left) and two b‐arches (right), forming short b‐arcades. Arrows denote b‐strands; and dotted lines denote H‐bonds. (B) Nomenclature used to describe residue positions in b‐arches. Open and filled circles denote side chains directed outside and inside the arch (o‐residues and i‐residues), respectively. Thick arrows denote b‐strands. Shaded region indicates the internal hydrophobic space of the arch. The number of A‐residues varies, depending on the arch type. (C) Simplified Ramachandran plot divided into seven discrete regions used to describe the conformations of b‐arch residues. The regions are superimposed on contours of ‘‘favored’’ regions that contain 98% of the non‐glycine and non‐proline residues of high resolution (1.8 A˚) protein structures (Lovell et al., 2003). Reprinted from Hennetin et al. (2006) with the permission of the publisher.

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bl

xbl

ab

bll

bed gbp

ppl

Two-residue arcs

Three-residue arcs

gbeb gbpl bepl

blbbl abebl

bllpbl

Four-residue arcs

Five-residue arcs

Six-residue arcs

Fig. 11. Pie diagrams showing observed percentages of different types of arc conformations in b‐solenoid proteins. Each circle corresponds to an arc with a certain number of residues. Areas of circles and their sectors are proportional to the number of observed cases. The arcade‐forming arcs are in bold and underlined. Reprinted from Hennetin et al. (2006) with the permission of the publisher.

short b‐strand. Considering large arcs in this way, it follows that they consist of two small arcs. For example, the most common five‐residue arc, blbbl, consists of two bl arcs separated by a buried b‐residue (Figs. 12 and 13). Similarly, the six‐residue bllpbl arc is a union of bll and bl arcs separated by a p‐residue. It is noteworthy that the standard arc conformations differ markedly from the recurring turn conformations, ag, pl, eg, ll, agl, and aagl, of b‐hairpins found in globular proteins (Efimov, 1993). The more residues there are in an arc, the greater the turn of the polypeptide chain (Fig. 12). Two‐residue arcs change the chain direction by 90–120 , three‐residue arcs by 90–180 , four‐residue arcs by 160–180 , and five‐ and six‐residue arcs by 180 . For two‐, three‐, and four‐residue arcs, the value of the turn‐angle depends on the size of the side chains of the b‐strands flanking the arc (in the 1B3 and 2B1 positions). Most of the relatively abundant b‐arches can form b‐arcades. An arcade is produced by several b‐arches with the same conformation stacked in‐register and stabilized by a network of peptide group H‐bonds (Fig. 13). Some b‐solenoid coils contain combinations of common arc conformations, and these arcs interact with each other via peptide group H‐bonds, as do stacked b‐strands. For example, one side of the B‐type coils of the C‐terminal domain of bacterial glutamate synthase (Binda et al., 2000; van den Heuvel et al., 2002) has several zigzagging b‐arcs that produce a segment that is, overall, linear and capable of forming inter‐coil H‐bonds. Some common relationships between b‐arches and their amino acid sequences are as follows: (i) although the interior positions of the b‐strands are mainly occupied by apolar (often Cb‐branched) residues, the 2B1

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position (Fig. 10B) frequently has polar residues—asparagine, threonine, serine, or cysteine—that form H‐bonds inside the arches (Fig. 12); (ii) the arc residues are predominantly polar or glycine; (iii) b‐arcs do not have as many proline residues as one might expect in view of the b‐strand‐breaking potential of this residue. Arches with the same conformation tend to have similar amino acid sequence patterns for key apolar, polar, or glycine residues (Hennetin et al., 2006). At the same time, the sequence patterns of the various kinds of arches differ in a characteristic manner (Fig. 12) and this information may be helpful for the prediction, modeling, and de novo design of b‐solenoids. It is noteworthy that the incidence of some b‐arcs correlates with the hand of the solenoid. For example, bl arcs are mostly found in right‐handed b‐solenoids. If the bl arc occurs in a left‐handed solenoid, its second position is frequently occupied by glycine. This is less common in right‐ handed solenoids. Conversely, ppl arcs are more frequent in left‐handed b‐solenoids. bll and blbbl arcs are observed only in right‐handed structures. Finally, L‐type b‐solenoids have an unusual inverted arch (Fig. 12). Stacking of these arches makes a groove which forms the center of the binding site for polysaccharides or pectins ( Jenkins and Pickersgill, 2001). In contradistinction to the other arches, the arc residues of inverted arches are interior‐facing and are apolar, while residues in the b‐conformation which bound the arc, face the solvent and are mostly polar (Fig. 12). These arcs have ab conformations. Frequently the first arc residue is small, glycine or alanine, and the second position is occupied by a bulky apolar residue.

D. Differences Between Sequences of Single‐Stranded and Triple‐Stranded b‐Solenoids As with single‐stranded b‐solenoids, TS ones also contain sequence repeats. However, whereas in the former, the repeat length usually matches the coil length (see above), in TS structures with their threefold symmetry, the sequence repeat length is typically one third of the coil length. Each repeat has one b‐strand with either the x or the xx sequence pattern. Unlike single‐standed b‐solenoids, where the interior ‐residues in positions close to turns are often alanine, glycine, serine, threonine, or asparagine, TS solenoids have bulky apolar residues in these positions. There are no asparagine ladders inside known TS b‐solenoids. The turns of the triangular coils of the TS b‐solenoid are made of three‐residue arcs with typical conformations gbp, axp, or axb (x has f 90 , c 140 ). The middle residue of the arc has a glycine‐specific conformation. Thus, the consensus sequence motif of these arcs is x‐Gly‐x (where x is any, but mostly a polar, residue).

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beb bed T

bll A

bl

bl N N

N

ppl N G

gbeb

gbpl G

bepl bebl

blbbl S

G ab

Fig. 12. A set of recurring b‐arches found in b‐solenoid proteins. In these schematized diagrams, the b‐strands are shortened and include only one residue from each of the b‐strands. Curved black arrows denote the polypeptide backbone. Blue, pink, and green circles show the locations of polar, apolar, and glycine side chains within the b‐arches, respectively. Open circles indicate positions that are not preferentially occupied by any particular type of residues. Letters inside some circles indicate certain amino acid residues which occur frequently (>30%) in particular positions. Italic letters describe b‐arc conformations (Fig. 10C). The b‐arches cluster into several groups, depending on the value of their turn‐angles: 90 in violet, 120 in blue, and 180 in orange. The five‐residue b‐arch can be represented by two 90 b‐arcs (red) and an inverted b‐arch is in green. The inset demonstrates the locations of these b‐arch modules within T‐, O‐, R‐, and L‐type b‐solenoids. Black linear modules indicate b‐strand extensions.

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b-Arcade

Fig. 13. Schematic diagram of a b‐arcade taken from the b‐solenoid domain of the yeast CAP (Dodatko et al., 2004). The arcade is formed by several b‐arches that contain five‐residue blbbl b‐arcs. The lower sidewall of the solenoid formed by b‐arcs is omitted to illustrate better the resemblance of this structure to the arcade. Reprinted from Hennetin et al. (2006) with the permission of the publisher.

The arcs most typical of TS b‐solenoids differ in conformation and consensus sequence from those of single‐stranded b‐solenoids (Hennetin et al., 2006). This difference may originate in the fact that each chain of a TS b‐solenoid has an axial rise of 14.5 A˚ as opposed to 4.8 A˚. On the other hand, the inverted arcs of the TS b‐solenoid from hyaluronidase HylP1 (Smith et al., 2005) have ab ‐conformations, like those found in single‐ stranded b‐solenoids (Hennetin et al., 2006). Neither the b‐strands nor the b‐arcs of TS b‐solenoids contain proline, while the long loops frequently contain this residue.

E. Examples and Perspectives of Sequence ‐Based Prediction of b‐Solenoid Structures While structural prediction of globular proteins remains uncertain in the absence of extensive sequence similarity to protein(s) of known structure (Moult, 2005), predictions can be quite reliable for proteins with repetitive substructures. Examples include collagen (Fraser and MacRae, 1973), a‐helical coiled coils (Cohen and Parry, 1994; Lupas, 1997), LRR proteins (Kajava, 2001; Kajava and Kobe, 2002), and b‐solenoids (Bateman et al., 1998; Kajava and Steven, 2006; Kajava et al., 2001; McDonnell et al., 2006). The efficacy of methods for predicting and modeling b‐solenoids, which have been comprehensively described, may be evaluated in systems where


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the molecular structure is subsequently determined experimentally. Recently, the crystal structure of a fragment of the FHA protein of B. pertussis was determined (Clantin et al., 2004). It provided strong confirmation of the b‐solenoid structure previously predicted (Kajava et al., 2001) for the domain that consists of an extended tract of 19‐residue sequence repeats called R1 repeats (Kajava et al., 2001; Makhov et al., 1994). The structural prediction was based on the following considerations. First, a b‐helical fold was inferred, based on CD spectroscopy and electron microscopy (Kajava et al., 2001; Makhov et al., 1994), as well as the a priori inference that repeats of about 20 residues are often b‐solenoids (Kobe and Kajava, 2000). Second, by analyzing an alignment of the sequence repeats, conserved—and presumably structurally important—sites were identified (Fig. 14). The distribution of conserved residues was recognized as a potential source of information about the disposition of strands and turns. Indeed, in known b‐helices, stretches of alternating conserved/variable positions correspond to strands (Jenkins and Pickersgill, 2001; Yoder and Jurnak, 1995); conversely, interruptions of this pattern should be turns. On this basis, three strands and three turns were identified in each repeat corresponding to one b‐helical coil of triangular shape (Fig. 14). Insertion/deletion sites detected in the repeat alignment were assigned to turns, consistent with the inferred arrangement. Then, selected fragments of known b‐helices were matched with this template to identify the most likely prototype for the 3D structure of a single coil. Finally, the coils were fused into a b‐helical structure which was refined by an energy minimization procedure. As with many confirmed b‐solenoids, the model was right‐handed with a shallow left‐handed twist. Comparison of this model with the corresponding part of the crystal structure (Fig. 14) shows their Ca‐traces to be virtually superposable, with an RMSD of 1.1 A˚ for the 2.5 repeats compared. Ongoing genome sequencing projects are revealing many proteins whose sequences contain b‐solenoid‐favoring repeats. For example, a recent survey of about 1000 virulence factors of Gram‐negative bacteria, which are secreted by the type V system via the autotransporter and two‐partner pathways, revealed that the majority have b‐solenoid domains ( Junker et al., 2006; Kajava and Steven, 2006). Structure prediction for such proteins promises to be a fertile research topic. Their high molecular weights and extended shapes are potential obstacles to crystallographic analysis, enhancing the potential impact of prediction. In turn, predictive models can provide a framework for mutational studies and facilitate the identification of fragments that may fold correctly and thus be suitable for study by X‐ray crystallography. These models should also facilitate solution of the corresponding crystal structures by molecular replacement (Rossmann and Arnold, 2001).

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A

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LxVxAGGxVxLxxLxAxGx b2

b1

b3

b2

b3

L V G

A

B

318

367

L A

V

L

Crystal structure G

G b1

397 443

Model

Fig. 14. Structural prediction and modeling of a fragment of FHA from B. pertussis containing R1‐repeats. (A) Successive stages in the modeling. From top to bottom: identification of the consensus sequence repeat, generation of 2D template of the coil, and the modeled 3D structure. In the consensus sequence, letters indicate residues that are conserved at the level of >60% identity. x is any residue; and filled circles represent bulky nonpolar residues. Apolar residues are in red; glycine in green. In the 2D template, open circles denote any (but mainly polar) residues, while filled circles denote conserved, mainly nonpolar, residues. Circles inside the coil contour indicate side chains located inside the structure and circles outside the contour denote side chains facing the solvent. Arrows indicate b‐strands. (B) A fragment of the crystal structure of FHA (Clantin et al., 2004) (on the top, in green color) and the 3D model (bottom, in brown).

F.

Prediction of Amyloidogenic Regions and Structures of Amyloid Fibrils

In view of the consideration that b‐solenoids and b‐arcades may also be structural elements of amyloid fibrils (Kajava et al., 2004; Lazo and Downing, 1998; Margittai and Langen, 2004; Pickersgill, 2003), sequence‐based detection and structure prediction of b‐solenoid proteins are pertinent to the identification of amyloidogenic sequences and the elucidation of amyloid fibril structures. As with b‐solenoid domains, amyloidogenic regions of


85

proteins may be enriched in glycine, Cb‐branched valine and threonine, as well as in asparagine and glutamine which are capable of forming H‐bonded ladders. At the same time, one can expect the occurrence of proline to be lower than average in amyloidogenic sequences. Furthermore, it is becoming apparent that amyloid fibrils formed by long (over about 30 residues) peptides are likely to have b‐arches stacked into b‐arcades (Antzutkin et al., 2003; Kajava et al., 2004; Ritter et al., 2005) similar to those found in b‐solenoids (Hennetin et al., 2006). On this basis, knowledge of the conformations and sequence motifs typical of b‐arches in b‐solenoids can be used to localize likely turn (arc) positions in amyloid fibrils and to model their structures.

VI.

Relationship Between b‐Solenoid Structures and Their Functions

A. Functional Implications of Elongated Shape and Rigidity of b‐Solenoids The majority of known b‐solenoid proteins are either virulence factors resident on the outer surfaces of bacteria or viral adhesins (Tables I and II; Fig. 3). They include enzymes that degrade or modify polysaccharides such as, for example, pectinases ( Johansson et al., 2002; Lietzke et al., 1996; Petersen et al., 1997; Pickersgill et al., 1998), adhesins (Clantin et al., 2004; Nummelin et al., 2004), and proteases (Baumann et al., 1993; Hege and Baumann, 2001; Otto et al., 2005). Bacteriophage tails that play a critical role in the early steps of infections frequently contain TS b‐solenoid domains (Kanamaru et al., 2002; Smith et al., 2005; Stummeyer et al., 2005; van Raaij et al., 2001). The high incidence of b‐solenoid domains in virulence factors implies that this structure is central to their function. We speculate that their elongated shapes allow them to protrude from the pathogen surface to interact with targets on host cells. A b‐solenoid may also be conducive to multivalent adhesion. Indeed, the large elongated surface provides enough space to accommodate multiple binding sites, while the apparent rigidity of a b‐solenoid affords a stable binding platform. This property may facilitate adhesion especially when it involves flexible molecules such as carbohydrates, which are known to be a common binding partner of b‐helices. Polysaccharide‐binding b‐solenoids often have a longitudinal groove formed by inverted b‐arches that represents a substrate‐binding site (Huang et al., 1999; Scavetta et al., 1999; Steinbacher et al., 1996). The majority of virulence factors also have tracts of irregular coils with long loops which, in some cases, fold into globular structures bulging from the b‐solenoid core (Emsley et al., 1996; Otto et al., 2005).

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These protruding structures may form catalytic sites or binding sites for small molecules. Remarkably, some polysaccharide modification enzymes of pathogenic bacteria are similar to enzymes of their host cells. For example, during cell development, pectin, a principal component in the primary cell wall of plants, is modified by its own pectin methylesterases that have b‐solenoid structures ( Johansson et al., 2002) and in this respect, resemble pectin lyases secreted by bacteria to break down these structures (Lietzke et al., 1996).

B.

Functional Implications of Highly Regular b‐Solenoid Structures

Insect antifreeze proteins (AFP) from the beetle T. molitor (Liou et al., 2000) and the spruce budworm C. fumiferana (Leinala et al., 2002) are perhaps the most regular protein structures yet observed. Despite differences in hand and coil shape (a right‐handed R‐type solenoid for the former and a left‐ handed T‐type solenoid for the latter), both solenoids are flat and have one conserved face formed by b‐strands with the Thr‐X‐Thr motif. The OH groups of the threonine side chains make a near‐perfect match with the ice lattice. These faces of the AFP solenoids represent a putative ice‐ binding surface, an element that is essential for AFP activity. Another example of a regular structure is the R‐type solenoid of MfpA protein from M. tuberculosis (Hegde et al., 2005). This protein binds to DNA gyrase and inhibits its activity. The regular solenoid structure exhibits size, shape, and electrostatic similarity to B‐form DNA. This DNA mimicry explains the inhibitory effect of this protein on DNA gyrase. These examples demonstrate that a highly constrained and regular b‐solenoid is an ideal platform for the design of regular two‐dimensional (2D) or helical structures.

C. b‐Solenoids as Oligomerization Motifs Several b‐solenoid domains appear to promote the oligomerization of multidomain proteins. There are at least three types of b‐solenoid association. First, oligomers (dimers or trimers) are formed by lateral interaction of the solenoids. For example, the C‐terminal domain of the bacterial cell division inhibitor MinC is a short right‐handed T‐type solenoid with an apolar lateral face that mediates homodimerization (Cordell et al., 2001). Trimers of several bacterial transferases are formed by lateral, in‐register, interaction of left‐handed T‐type b‐solenoids (Fig. 5). Second, dimers may form via interactions of the ‘‘open’’ terminal coils of b‐solenoids as in the dimeric structure of iron transporter stabilizer SufD (Badger et al., 2005). Finally, dimerization may be mediated by swapping of b‐strands of the terminal coils, as in the CAP (Dodatko et al., 2004) (Fig. 3).


87

Unlike other oligomerization modules such as, for example, a‐helical coiled coils (Burkhard et al., 2001), the oligomerization of b‐solenoids has also an additional role, that is, the creation of grooves, pockets, or concave surfaces in regions of intermolecular contacts to serve as binding sites for other molecules (Fig. 5). For example, triangular b‐solenoids of bacterial transferases pack side‐by‐side, in‐register, to form three identical, very narrow, clefts along the interfaces between subunits (Figs. 3 and 5). Crystallographic studies have shown that these clefts can bind a variety of small molecules such as acetyl‐coenzyme A (Sulzenbacher et al., 2001), Zn2þ (Kisker et al., 1996), pimelate‐succinyl–CoA complex (Beaman et al., 2002), antibiotic dalfopristin (Kehoe et al., 2003), and cysteine (Pye et al., 2004). The clefts form the active sites of these enzymes. Another example is a homodimer of the C‐terminal actin‐binding domain of eukaryotic CAP. Dimerization of the b‐solenoid generates a ‘‘corner’’ (Fig. 3) with a concave surface that may be a binding site for either G‐actin or N‐terminal domains of CAP (Dodatko et al., 2004).

D. b‐Solenoids as Scaffolds for Multidomain Complexes The rigidity and elongated shape of b‐solenoids can be used to effect the precise positioning of globular domains in multidomain complexes. For example, an ammonia tunnel involved in signal transduction among redox and active centers of bacterial glutamate synthase is formed at the point where three globular domains meet. This tripartite structure is stabilized by a b‐solenoid domain (Binda et al., 2000; van den Heuvel et al., 2003). The dimeric structure of iron transporter stabilizer SufD (Badger et al., 2005) may also have a role in positioning functional domains relative to each other. The crystal structure of potato tuber ADP‐glucose pyrophosphorylase ( Jin et al., 2005) revealed that its b‐solenoid is involved in cooperative allosteric regulation and assembly of the enzymatic tetramer. Remarkably, the left‐handed T‐type structure of the ADP‐glucose pyrophosphorylase b‐solenoid resembles the b‐solenoids that trimerize bacterial transferases. However, the pyrophosphorylase solenoid does not form laterally associated oligomers.

VII.

Evolution of b‐Solenoid Proteins

A. Independent Evolutionary Paths of b‐Solenoid Folds Despite a common topology and, in some cases, similar functions, it appears likely that many b‐solenoid domains emerged independently

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rather than descending from a common ancestor. Comparison of the 38 known b‐solenoids of the nonredundant set (Tables I and II) suggests that they partition into 15 subsets that may derive from independent evolutionary paths. The most remarkable cases are non‐homologous b‐solenoid proteins that execute the same function. For example, structurally distinct antifreeze proteins (a right‐handed R‐type solenoid for the AFP from T. molitor (Liou et al., 2000) and a left‐handed T‐type solenoid for the AFP from C. fumiferana (Leinala et al., 2002)) have almost identical ice‐like surfaces formed by b‐strands with the Thr‐X‐Thr motif. Apparently, this functionally important surface emerged more than once as a result of convergent evolution. Another example is given by adhesins of pathogenic Gram‐negative bacteria. Despite similar functions, their b‐solenoids are very different: FHA from B. pertussis has a right‐handed T‐type solenoid (Clantin et al., 2004; Kajava et al., 2001); P.69 pertactin from B. pertussis contains a right‐handed L‐type solenoid (Emsley et al., 1996); and YadA adhesin from Yersinia enterocolitica has a trimeric left‐handed O‐type solenoid (Nummelin et al., 2004). Recent bioinformatic analysis suggested many other examples of virulence factors from bacterial pathogens with similar shapes and functions but different solenoid structures (Kajava and Steven, 2006). Finally, the TS b‐solenoids of phage tail proteins, being very different from each other, may represent another example of convergent evolution. The inferred tendency of b‐solenoids to evolve independently can be explained by the repetitive character of their sequences. Repetitive sequences can evolve more rapidly than nonrepetitive ones (Buard and Vergnaud, 1994; Marcotte et al., 1999). Therefore, a gene for a b‐solenoid protein can be created by multiple duplications of the DNA coding for a short ancestral peptide. If the ancestral peptide had a predisposition to polymerize into a stack of coils linked by b‐structural H‐bonds, this property could develop into a b‐solenoid folding pathway for a multicoil descendent. (One might call the proposed mechanism ‘‘one repeat‐one solenoid.’’) A similar evolutionary mechanism has been suggested for other solenoid proteins (Kajava, 1998; Kobe and Kajava, 2000). Analysis of known b‐solenoids supports this scenario. Repeats of the same type typically occur in tandem blocks. When different types of repeats coexist in one protein, they cluster in separate tandems (e.g., in FHA from B. pertussis and lspA1 protein from H. ducreyi) (Kajava and Steven, 2006; Kajava et al., 2001). An additional reason for this coherence is that coils of different types do not pack together well and cannot accomplish the specific inter‐repeat H‐bonding networks that occur between adjacent coils of the same type. Therefore, transformation of one type of tandem repeat to another may be hampered by the need for simultaneous mutations in several repeats.


89

Thus, both the potential of sequence repeats to evolve quickly and selective pressure from the structure may favor an evolutionary scenario in which some b‐solenoids emerged independently. We note that sequence repeats may be deleted as quickly as they are inserted. Thus, repetitive sequences are well suited to respond rapidly to environmental changes (Buard and Vergnaud, 1994). This property may contribute to the abundance of b‐solenoids at bacterial and viral surfaces.

B.

Homologous b‐Solenoid Proteins

Although b‐solenoids may have evolved along multiple pathways, they also form families of homologous proteins. Jenkins and coauthors (1998) have suggested that the right‐handed L‐type b‐solenoids constitute a single family descended from a common ancestor. This family contains enzymes that degrade and/or modify polysaccharides. Among them are bacterial, plant, and fungal proteins, suggesting that bacteria acquired these b‐solenoid proteins once by horizontal gene transfer, probably from host plant or fungal DNA. In the current nonredundant set of b‐solenoids with low (7‐M urea in addition to normal SDS sample buffer and heating to 100 C to reproducibly exhibit the Ure2p

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band on Western blots (Baxa et al., 2003; Ripaud et al., 2004; Schlumpberger et al., 2000). In consideration of thermal stability, a calorimetric investigation of Ure2p prion domain‐containing filaments detected no evidence of these domains denaturing up to 105 C (Baxa et al., 2004). In comparison, most proteins denature at temperatures of 50–70 C and rarely exceed 80–90 C, except for proteins of extreme thermophiles. Sup35p filaments resist dissociation by SDS unless they are heated, and assays based on this effect (dissociation temperature in 1.6% SDS) have been used to distinguish different types of filaments (Chien et al., 2003; Tanaka et al., 2004).

D. Demonstration of Cross‐b Structure Cross‐b structure has been demonstrated for Sup35pNM filaments. Serio et al. (2000) observed a 0.47‐nm reflection by X‐ray diffraction, and subsequently this reflection was shown to be meridional both by X‐ray fiber diffraction (Kishimoto et al., 2004) and electron diffraction (King and Diaz‐ Avalos, 2004). In the Ure2p system, cross‐b structure has been established by electron diffraction from prion domain filaments preserved in vitreous ice (Fig. 7; Baxa et al., 2005). In addition, a 0.47‐nm reflection was detected by both X‐ray diffraction and electron diffraction from filament preparations of full‐length Ure2p and the Ure2p1–65‐GFP fusion, indicating that they contain the same structure (Fig. 7; Baxa et al., 2005). Analysis of HET‐s218–289 filaments by solid‐state NMR yielded evidence for b‐strand conformation (Ritter et al., 2005). Melki and coworkers currently maintain that Ure2p filaments are not amyloid (Bousset et al., 2002), based on their failure to observe the 0.47‐nm reflection indicative of cross‐b structure by X‐ray diffraction (Bousset et al., 2003). We have observed a strong reflection at this spacing by both X‐ray diffraction and electron diffraction, both for Ure2p filaments and for prion domain filaments but not for C‐domain preparations (Baxa et al., 2005). Recently, the same group (Fay et al., 2005) attributed these results to an artifact of drying but apparently failed to notice or to appreciate the significance of the fact that the reflection was also detected for filaments preserved in their native states in vitreous ice.

E. Natively Unfolded Conformations In recent years it has become apparent that many proteins have substantial regions (>30 amino acids) that lack a definite fold (Dunker et al., 2002; Fink, 2005; Tompa, 2002; Uversky, 2002). Such regions are called

STRUCTURAL BIOLOGY OF FUNGAL PRIONS

147

Fig. 7. Electron diffraction of Ure2p filaments. (A) Diffractograms of sucrose‐embedded specimens of Ure2p10–39 (left upper quadrant), Ure2p (left lower), Ure2p1–65 (right upper), and Ure2p1–89 (right lower). The 0.47‐nm reflection indicative of cross‐b structure is indicated. (B) Diffractograms of vitrified (frozen, hydrated) specimens of Ure2p (left) and Ure2p10–39 (right). The 0.47‐nm reflection from cross‐b structure is marked as is the sharp 0.37‐nm reflection from small crystals of cubic ice that condensed as contaminants on this specimen during cryo‐transfer into the electron microscope. The broad reflections at 0.37 nm and 0.22 nm from diffraction from the vitreous ice layer (Dubochet et al., 1982) in which the filaments were embedded are also present (the part of the spectrum containing latter reflection is not shown). (C) Azimuthally averaged, baseline‐corrected scans of the region around the 0.47‐nm reflection of the vitrified specimens: Ure2p filaments—gray; Ure2p10–39 filaments—black. All panels adapted from Figures 1 and 2 of Baxa et al. (2005).

natively unfolded or intrinsically unstructured and are characterized by extreme sensitivity to proteases and by random coil signals when observed by spectroscopic methods such as CD or NMR. Many of them convert to folded states when they bind to specific interaction partners and such conversions have been implicated in the regulation of diverse processes (Tompa, 2005). NMR spectroscopy of purified soluble Ure2p has shown that an N‐terminal tract of about 90 amino acids is unfolded (Pierce et al., 2005). This observation explains the acute sensitivity to proteolysis of the N‐domains of soluble Ure2p (Baxa et al., 2003; Thual et al., 1999). Similarly, NMR analysis of soluble HET‐s showed the region of amino acids 218–289 to be unfolded (Balguerie et al., 2003). Circular dichroism measurements on soluble Sup35pN and NM domains indicate a largely unfolded structure (Glover et al., 1997; King et al., 1997). There are as yet no such data for soluble Rnq1p, but the algorithms FoldIndex, Globplot, and RONN (Linding et al., 2003; Prilusky et al., 2005; Yang et al., 2005) which correctly predict an unfolded conformation for Ure2p1–89 and Sup35p1–114 make similar predictions for Rnq1p153–405 (U.B., unpublished results).

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As noted above, many natively unfolded proteins become folded in the presence of an appropriate interaction partner. Fungal prion domains subscribe to this paradigm whereby the partner is itself (each other) and the interaction represents homotypic polymerization into amyloid.

F. Functional Domain of Ure2p Retains Its Native Fold in Filaments An important question with respect to modulation of activity is whether the functional domains change their structures on entering the filamentous state. Several lines of evidence indicate that such is not the case for Ure2p. Fusion proteins of its prion domain with various enzymes all form filaments in which the appended proteins remain active, indicating that their conformations are unchanged (Baxa et al., 2002). Although the GST‐ like functional domain of Ure2p does not transfer glutathione to any known substrate, it binds glutathione (Bousset et al., 2001b) and it has glutathione peroxidase activity (Bai et al., 2004). Both activities are retained in Ure2p filaments (Bai et al., 2004; Bousset et al., 2002), suggesting that the C‐domain remains folded. Further support for this conclusion was obtained by scanning calorimetry, whereby it was demonstrated that the thermal denaturation profile of the C‐domain was unchanged whether the domain was soluble or in filaments (Baxa et al., 2004). A similar scenario is assumed for Sup35p (Chien et al., 2004; Glover et al., 1997). The inactivation of Ure2p and by inference also of Sup35p in the prion states comes from steric constraints. One such mechanism would be steric blocking, that is, the binding sites for the interaction partners are not accessible in filaments.

G. Conformational Changes Accompanying Filament Formation For proteins in which conformational changes between two states are suspected but for which high resolution structural information is hard to come by, spectroscopic methods—circular dichroism, FT‐IR, and Raman spectroscopy—offer an approach. The spectra may be analyzed to obtain estimates of secondary structure content for both states. The ability to demonstrate a conformational change depends both on the size of the change and the margin of error in the estimates. These methods have been applied quite extensively to prion proteins (Table III). The secondary structure of Ure2p both in filaments and in the soluble state are now quite well defined by other methods: a crystal structure of the functional domain (75% of the protein) has been determined; its conformation is unchanged in filaments; in soluble Ure2p, the prion domain is random coil, and in filaments, it is amyloid. Accordingly, it is potentially informative

Table III Secondary Structure Estimates by Spectroscopic Methods in Prion Systems Specimen Ure2p Ure2p95–354

Crystal

References

Constructa

Alpha % (res)

Beta % (res)

Turn/coil % (res)

Bousset et al. (2001a), Umland et al. (2001) Data cited in Baxa et al. (2003) Bousset et al. (2003) Bousset et al. (2002)

47 (123)

5–6 (16)

47 (123)

49 (134) 33 (86) 63 (164)

16–21 (52) 21 (55) 22 (57)

30–35 (89) 46 (120) 15 (36)

35 (123) 36 (127) 45 (159) 60 (212)

5 (18) 30 (106) 30 (106) 17 (60)

60 (213) 34 (120) 25 (88) 23 (81)

35 (123)

16 (56)

49 (175)

14 (50) 32 (113) 29 (103) 74 (262) 40 (142)

49 (173) 40 (142) 40 (142) 20 (71) 28 (99)

37 (131) 28 (99) 31 (110) 6 (21) 31 (110)

0 (0) 14 (11)

0 (0) 67 (54)

100 (90) 20 (16)

51 (48) 0 (0) 0 (0)

3 (3) 0 (0) 0 (0)

49 (45) 100 (65) 100 (94)

0 (0) 14–24 (13)

57 (37) 45–67 (40)

43 (28) 19–32 (18)

Ure2p Ure2p95–354 crystals Ure2p95–354 soluble

Raman FT‐IR FT‐IR

Ure2p soluble Ure2p soluble

Pierce et al. (2005) Taylor et al. (1999)

Ure2p soluble

NMR/crystal Raman I Raman II FT‐IR

Ure2p filaments

Model/crystal

Ure2p filaments Ure2p filaments Ure2p filaments Ure2p filaments

Raman I Raman II FT‐IR FT‐IR FT‐IR

Bousset et al. (2001a), Kajava et al. (2004), Umland et al. (2001) Taylor et al. (1999)

Ure2p1–90 solubleb Ure2p1–80 solubleb

NMR Calc difference

Ure2p1–94 solubleb Ure2p1–65 solubleb Ure2p1–94 solubleb

Calc difference CD CD

Pierce et al. (2005) Data cited in Baxa et al. (2003), Taylor et al. (1999) Bousset et al. (2002) Baxa et al. (2002) Thual et al. (2001)

Ure2p1–65 filaments Ure2p1–69 filaments

Model FT‐IR

Kajava et al. (2004) Schlumpberger et al. (2000)

Bousset et al. (2002)

Schlumpberger et al. (2000) Bousset et al. (2002) Bousset et al. (2003)

149

(continued)


80–354

Method

Table III (continued) Method

References

Constructa

Alpha % (res)

Beta % (res)

Turn/coil % (res)

Raman I Raman II

Taylor et al. (1999)

Sup35p Sup35p2–114 soluble Sup35p2–114 filament Sup35p1–253 soluble Sup35p1–253 filament

CD CD CD CD

King et al. (1997) King et al. (1997) Glover et al. (1997) Glover et al. (1997)

HET‐s HET‐s soluble HET‐s soluble HET‐s filaments HET‐s filaments

FT‐IR CD FT‐IR CD

Dos Reis et al. (2002) Dos Reis et al. (2002) Dos Reis et al. (2002) Dos Reis et al. (2002)

30 (87) 34 (98) 10 (29) 17 (49)

20 (58) 16 (46) 45 (130) 32 (92)

50 (144) 50 (144) 45 (130) 50 (144)

PrP PrPC full

NMR

30 (62)

4 (8)

66 (138)

PrPC full (hamster) PrPC full (hamster) PrP27–30 PrP27–30 PrP27–30 PrP27–30 PrP27–30

FT‐IR CD FT‐IR CD FT‐IR FT‐IR FT‐IR

Lopez Garcia et al. (2000), Zahn et al. (2000) Pan et al. (1993) Pan et al. (1993) Caughey et al. (1991) Safar et al. (1993) Wille et al. (1996) Pan et al. (1993) Gasset et al. (1993)

42 (88) 36 (75) 17 (24) 0 (0) 23 (33) 21 (30) 25 (35)

3 (6) – 47 (67) 43 (61) 48 (68) 54 (77) 54 (77)

55 (115) – 36 (51) 57 (81) 29 (41) 25 (35) 21 (30)

PrPSc PrPSc PrPSc

Model CD FT‐IR

Govaerts et al. (2004) Safar et al. (1993) Pan et al. (1993)

23 (47) 20 (42) 30 (63)

23 (48) 34 (71) 43 (90)

54 (113) 46 (96) 27 (56)

Ure2p1–65 filaments

65 (42) 60 (39)

35 (23) 32 (21)

Mostly random coil Mostly b‐sheet Mostly random coil Mostly b‐sheet

Block diagrams in the column are used to indicate the length of the protein construct used in the experiment (gray area) compared to the whole protein (full rectangle). b In the context of the full‐length Ure2p protein or in fusion proteins of Ure2p1–65 with different enzymes. Rows that are calculated based on structures (experimental crystal or NMR structures or models) are marked in bold.

BAXA ET AL.

a

0 (0) 8 (5)

150

Specimen


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about the scope of spectroscopic data pertaining to other prion proteins to review the Ure2p data. With amyloidogenic proteins, one expects an enhancement of b‐structure as they enter the amyloid state. For prion domain filaments, a high b‐sheet content was detected both by Raman spectroscopy (Taylor et al., 1999) and FT‐IR (Schlumpberger et al., 2000), consistent with its being amyloid by other criteria. Although this domain does not remain in solution to allow a measurement in that state, NMR analysis has shown that this domain is unfolded in the context of soluble full‐length Ure2p (Pierce et al., 2005). For spectroscopic studies of full‐length Ure2p, the complication arises that the amyloid‐forming component constitutes only 15–25% of the molecule, depending on how far the prion domain extends (Section II.A). The functional domain is mostly a‐helix with little b‐sheet. Therefore, one should expect the b‐sheet content to be initially low and to increase by 5–10% in filaments and the a‐helix content to be unchanged. However, the data show some considerable discrepancies: (1) FT‐IR data for Ure2p filaments have yielded estimates of a‐helix content as high as 74% (Bousset et al., 2003), the theoretical value being 35%. (2) Two FT‐IR analyses of the functional domain by the same group diverge by as much as 30% in a‐helix (one being 15% higher and the other 15% lower than the crystal structure (Bousset et al., 2002, 2003)). (3) Our estimates of the functional domain by Raman spectroscopy are off by as much as 15% in b‐sheet (data cited in Baxa et al., 2003). Constructs of Sup35p (Sup35pN and Sup35pNM) have been investigated by CD spectroscopy and in both cases an increase of b‐structure on filament formation was reported (Glover et al., 1997; King et al., 1997). No attempt was made to quantitate secondary structure.

V.

Experimentally Derived Constraints on Prion Filament Structure

We now summarize current experimentally derived constraints that models of yeast prion filaments must satisfy in addition to the basic requirement of cross‐b structure and then go on in Section VI to discuss their implications for several models that have been proposed.

A.

Polarity

A basic structural property of protein filaments is polarity, that is, directionality. Almost all naturally occurring filaments are polar (e.g., F‐actin, microtubules, TMV, and so on). The few exceptions are either bipolar, like myosin (Huxley, 1963; Squire, 1981), or nonpolar, like intermediate filaments (Herrmann and Aebi, 2004). One method of determining

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Fig. 8. Polarity and handedness of prion filaments. (A, B) Atomic force microscopy (AFM) study of Sup35p filament growth. (A) Image of a Sup35pNM seed with tag, labeled with antibody (thick part), and elongated by addition of untagged Sup35pNM monomers (thin part). The AFM signal (height above substrate) is color coded as shown. Bar ¼ 200 nm. (B) Statistics of the growth patterns of 200 individual filaments. Notice that the large majority of filaments show asymmetric growth, that is, long side short side, indicative of polar structures. Panels (A) and (B) were reprinted from DePace and Weissman (2002) with copyright permission from Nature Publishing Group. (C) Polar growth of Sup35pNM filaments demonstrated by fluorescence microscopy. Red‐labeled Sup35pNM filaments were elongated by the addition of green‐labeled Sup35pNM monomers. Panel (C) was reprinted from Figure 1 of Inoue et al. (2001) with copyright permission from the American Society of Biochemistry and Molecular Biology. Bar ¼ 2000 nm (D, E) Unidirectional shadowing of Ure2p1–80‐GFP shows a long‐pitch left‐ handed twist (indexed by white arrows). Deposited metal is white and shadows are black. Panel (E) shows the internal standard, bacteriophage T4 polyheads whose low‐pitch helices (indexed in white lines) are known to be right‐handed. Black arrows indicate the shadowing direction. Bar ¼ 50 nm. Panels (D) and (E) adapted from Figure 2 in Kajava et al. (2004). (F) AFM of Ure2p filaments ( Jiang et al., 2004). Note the orientation


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whether a filament is polar or nonpolar is to measure the growth rates of the two ends separately. If these rates differ, the filament should be polar; on the other hand, if similar growth rates are observed, this may mean either that the filament is nonpolar or that the two ends coincidentally have similar growth rates. In such cases, further information is needed. Observations of this kind may be performed by using two batches of subunits that may be distinguished by different labels. In the Sup35p system, one study used AFM to monitor the growth of filaments from two constructs, one of which had a hemagglutinin epitope that could be recognized by antibodies (DePace and Weissman, 2002; Fig. 8). Filaments having the epitope were used to seed solutions of the other construct, so that thin parts grow out of the thick seed (antibody bound). These results clearly indicated that Sup35p filaments are polar (Fig. 8). Interestingly, different classes of growth rates could be distinguished suggesting the existence of structural variants (DePace and Weissman, 2002)—see Sections VII.B and VIII.B. Similar results have been obtained with gold staining of Sup35pNM filaments (Scheibel et al., 2001) and with fluorescence labeling (Inoue et al., 2001; Fig. 8). Fluorescence microscopy of labeled filaments was also used to investigate the polarity of Ure2p filaments (Fay et al., 2003). Very similar behavior as with Sup35p filaments was observed. Most filaments were found to grow only at one end; only few grew at both ends and these showed different growth rates at the two ends. Taken together, these observations leave little doubt that both Sup35p and Ure2p filaments are polar with the possibility to grow on both ends (bidirectional) as has also been observed for other amyloids such as amylin (Goldsbury et al., 1999).

B.

Handedness

Each kind of polar filament has an intrinsic handedness. Ure2p1–80‐GFP filaments have been determined to have a left‐handed twist by electron microscopy of specimens contrasted by unidirectional heavy metal shadowing (Kajava et al., 2004; Fig. 8). Atomic force microscopy also images the upper surface of the specimen and can therefore be used to reveal the hand of a filament. AFM images of Ure2p filaments (Jiang et al., 2004) also indicate a left‐handed twist (Fig. 8). Since b‐sheets generally have left‐ handed twists, these observations are consistent with the b‐sheets in these amyloid structures being aligned with the filament axes. of the high regions of the filaments that suggest a left‐handed twist. Bar ¼ 200 nm. Panel (F) was reprinted from Figure 6 in Jiang et al. (2004) with copyright permission from the American Society of Biochemistry and Molecular Biology.

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Fig. 9. Scanning transmission electron microscopy of Ure2p filaments. (A, B) STEM images of (A) Ure2p and (B) Ure2p1–80‐GFP, negatively stained with vanadate. White arrows mark the 4‐nm core fibrils. The black asterisk indicates a reference TMV particle (Bar ¼ 50 nm). (C) Example of STEM dark‐field image of unstained freeze‐dried Ure2p filaments used for mass‐per‐unit‐length analysis (Bar ¼ 100 nm). (D) Histogram of mass‐ per‐unit‐length measurements from Ure2p filaments. The measurements were fitted with a Gaussian distribution (solid curve). (E) Graph summarizing mass‐per‐unit‐length


C.

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Mass‐Per ‐Unit‐Length

The parameter of mass‐per‐unit‐length, if known, places a strong constraint on models for the packing of subunits in filaments. Scanning transmission electron microscopy (STEM) (Wall and Hainfeld, 1986) has been used to measure this parameter for filaments of various Ure2p constructs, including full‐length Ure2p, Ure2p1–65, and several fusion proteins (Fig. 9; Baxa et al., 2003). For all specimens, the data yielded a value of about one subunit per 0.47‐nm axial rise. Extensive measurements of this kind have also been made for Sup35p1–61‐GFP (Diaz‐Avalos et al., 2005). In that study, filaments that induce three different variants (Section VIII.B) were classified into four morphological kinds. Small differences between their distributions of mass‐per‐unit‐length data were reported whose basis is still under discussion (Diaz‐Avalos et al., 2005): however, in all cases, the values obtained were within 20% of one subunit per 0.47 nm.

D. EM Images of Filament Structure In the absence of high‐resolution data from X‐ray crystallography or solution NMR, EM has been a primary source of information about prion filament architecture. In particular, STEM micrographs of specimens stained with vanadate have yielded some very informative images (Fig. 9; Baxa et al., 2003; Diaz‐Avalos et al., 2005; Ksiezak‐Reding and Wall, 2005). Micrographs of Ure2p filaments show a core fibril of about 4‐nm diameter surrounded by globular domains that appear to be loosely packed and not highly ordered. A similar impression is conveyed by cryo‐EM images of Ure2p filaments embedded in vitreous ice (Baxa et al., 2003). The vanadate‐stained STEM data further indicate that, at a given axial level in a filament, the globular domains are not symmetrically distributed about the core fibril but are clumped to one side and this distribution precesses around the axis, giving the filament a sinusoidal or corkscrew appearance, even though the core fibril is more or less straight. This effect is particularly evident for Ure2p N‐domain/GFP fusions (Section VII.B). An electron micrograph of negatively stained filaments of full‐length Sup35p suggests a backbone surrounded by peripheral material (Glover et al., 1997). for numerous Ure2p constructs. Filled data points are averaged measurements from single filaments for a given construct; open points are halved measurements from paired filaments: Ure2p—red; Ure2p1–65—gray; Ure2p1–65‐barnase—blue; Ure2p1–65‐carbonic anhydrase—yellow; Ure2p1–65‐GST—green; Ure2p1–65‐GFP—orange; Ure2p1–80‐GFP— light green. The domain structures of these constructs are shown schematically at right, with the prion domain in gray and the appended domain colored as in the data points. Two versions of the prion were used; residues 1–65 and 1–89. The latter includes most of the linker region (darker gray). All panels adapted from Figures 5 and 6 of Baxa et al. (2003).

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E.

Parallel In‐Register Arrangements of b‐Strands

Strands in b‐sheets can have either a parallel or an antiparallel arrangement (see Kajava et al., this volume; Pauling and Corey, 1951). In globular proteins, antiparallel b‐sheets predominate; in fact, extended parallel b‐sheets were only observed in connection with a‐helical structures (the so‐called a/b‐proteins) until purely parallel b‐sheets were found in b‐helical proteins (Kajava and Steven, this volume; Kobe and Kajava, 2000; Yoder et al., 1993). Recently, two experimental procedures have emerged as capable of distinguishing between parallel and antiparallel b‐sheet conformations in amyloids—solid‐state NMR spectrosopy (Antzutkin et al., 2000; Benzinger et al., 1998; Lynn and Meredith, 2000; Tycko, 2000) and ESR spectroscopy (Der‐Sarkissian et al., 2003; Jayasinghe and Langen, 2004; Torok et al., 2002). These studies have shown that the b‐sheets present in amyloids of a‐synuclein, islet amyloid polypeptide (Der‐Sarkissian et al., 2003; Jayasinghe and Langen, 2004), and Ab peptide (Antzutkin et al., 2000; Torok et al., 2002) are parallel. NMR analysis of Ure2p10–39 filaments detected parallel b‐sheets and provided evidence for an in‐register arrangement of the polypeptide chains, that is, successive residues along one chain are close to the corresponding residues of a neighboring chain. It was further shown that Gln side chains form stacks in which the side chains can H‐bond to each other (Chan et al., 2005). Other experiments with Cys mutants and fluorescence labels and chemical cross‐linking on Sup35pNM filaments (Krishnan and Lindquist, 2005) support the view that the regions from residues 21 to 38 and from 86 to 106 may also have a parallel in‐register arrangement.

F.

Filament Formation Depends Mostly on Amino Acid Composition, Not Sequence

In general, protein folds depend on and indeed are specified by amino acid sequence: in many instances, small changes are sufficient to destabilize a fold or to render a sequence incompetent to fold. In this context, it is striking that mutants in which the prion domain sequences of Ure2p (Ross et al., 2004) and Sup35p (Ross et al., 2005) were radically altered by random permutation retained their prionogenic character and, in the case of Ure2p at least, the ability to form filaments. For Sup35p, it has also been reported that a large part of the N‐terminal domain can be exchanged with poly‐Gln without changing the ability to form prions (DePace et al., 1998). These observations indicate that the ability of these proteins to propagate as prions resides mostly in the amino acid compositions of their prion domains. Credible filament models should be able to account for this unusual property.


VI.

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Structural Models for Prion Amyloid Filaments

In this section, we discuss several recent proposals for how the proteins are arranged in amyloid filaments of fungal prion proteins in light of current experimental data.

A. Superpleated b‐Structure In this model, also called the b‐serpentine, each polypeptide chain zig‐zags in a planar serpentine fold and successive chains are stacked in‐ register, one on top of another (Kajava et al., 2004; Fig. 10). This arrangement generates an array of parallel b‐sheets, each composed of identical strands and aligned with the filament axis. The strands run perpendicular to the filament axis, that is, it is a cross‐b structure. The strands in a given b‐sheet are oriented in the opposite direction to those in the next adjacent b‐sheet. In the regions between adjacent b‐sheets, called ‘‘bays,’’ adjacent sheets interact via the side chains of apposed strand residues. The bays may be occupied by polar uncharged residues, for example, Asn and Gln can form H‐bonded ladders, or by hydrophobic residues. Charged residues are segregated to turn regions where they have the greatest room for maneuver so as to minimize the energetic penalty arising from electrostatic repulsion of juxtaposed residues of like charge. This model is a polar cross‐b structure with a left‐handed twist that complies with the mass‐per‐unit‐length data: moreover, it readily accommodates sequence randomization because like residues are still stacked over like residues, regardless of their order, and sequence permutation does not increase the number of charged residues or prolines which would be most likely to destabilize structures of this kind (Fig. 10). The configuration of strands and turns allows some variation without putting charged residues inside the core structure. We envision that structural variations of this kind offer a plausible explanation for the phenomenon of prion variants, as discussed in Section VIII. As proposed (Kajava et al., 2004), the model of the complete Ure2p prion domain has nine b‐sheets with a predicted core fibril cross section of 3‐nm wide by 7‐nm long. These dimensions are not necessarily at odds with the reported estimate of 4 nm for the average width (Baxa et al., 2003; Taylor et al., 1999; Section III.C) since molecular dimensions in this range are difficult to measure accurately from negatively stained specimens and are further complicated in this instance by the tendency of core fibrils to bundle and, in images of intact Ure2p filaments, they are largely obscured by the larger C‐domains. Nevertheless, the model is adaptable while maintaining the same basic principle by peeling off one or more sheets from the end: this would give a smaller length dimension. It has also been shown

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that the prion domain fragment, Ure2p10–39 forms similar amyloid fibrils (Chan et al., 2005; Kajava et al., 2004). The model readily accommodates this observation in terms of a filament with a reduced number of sheets—four, in this case, Fig. 10—and predicts that other fragments will share this property. Originally formulated for Ure2p, it has been proposed that models of this kind are also applicable to many other kinds of amyloids including filaments of Sup35p, polyglutamine and a‐synuclein (Kajava et al., 2004), and amylin (Kajava et al., 2005). Less detailed models with similar topologies have been considered for amyloid filaments of a‐synuclein (Der‐Sarkissian et al., 2003), tau protein (Margittai and Langen, 2004), and streptococcal protein G (Wang et al., 2005).

B.

b‐Helical Models

The b‐helix is a well‐authenticated fold, having been observed in more than 20 crystal structures, mostly of secreted bacterial proteins ( Jenkins and Pickersgill, 2001; Yoder and Jurnak, 1995; see also Kajava and Steven, this volume). In a b‐helix, the polypeptide chain winds around the molecular axis, each coil consisting of three short b‐strands with connecting turns (Fig. 11). Corresponding strands in successive turns are stacked, generating narrow parallel b‐sheets that are aligned with the Fig. 10. The super‐pleated b‐structure model. (A) Diagram of the b‐serpentine fold proposed for the Ure2p prion domain in the superpleated b‐structure model of the filament (Kajava et al., 2004). As shown here, the model has 9 b‐strands but it is adaptable to fewer (loc cit.); for instance, the subdomain constituted by residues 10–39 forms filaments and may be described as a four‐sheet portion of the model. The prion domain has few charged residues (circled): the only 4 charged residues in the first 64 residues are spaced regularly, 7 residues apart, and the same spacing separates the next 2 charged residues, Arg‐65 and Asp‐72. (B) Ball‐and‐stick representation of the four‐strand serpentine model for Ure2p10–39 produced by an energy‐minimization calculation (Kajava et al., 2004). The stucture is stabilized by a network of H‐bonds (red broken lines). Water is expected to be largely excluded from the regions (bays) between adjacent b‐sheets by close packing of polar uncharged and apolar side chains, as illustrated by the gray dotted van der Waals contour surrounding the polypetide chain. The backbone is in purple. Carbon, oxygen, and nitrogen atoms are in green, red, and blue, respectively. The dispositions of the side chains of the charged residues, in turn positions, may vary at successive levels in the stack in order to accommodate their mutual electrostatic repulsion. Panels (A) and (B) adapted from Figures 1 and 3 of Kajava et al. (2004). (C) Schematic presentation of different structural variants of the super‐pleated b‐structure model. Envisaged variations on this theme involve turns enlarging into loops of different sizes, and strands of different lengths, or serpentines folded from different segments of the full prion domain. Moreover, it could also include different conformations of loops and side chains (not shown).

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Fig. 11. b‐helical filament models. (A) Ribbon diagram representation of four coils of a generic right‐handed b‐helix. The three b‐strands in a given coil are colored red, blue, and green. (B) Schematic diagram of a b‐solenoid, showing three axially stacked subunits with two coils per subunit. Here the color coding denotes different subunits (red, black, gray), not different strands in the same subunit, and strands are represented simply as straight segments. A model of this kind—specifically, one involving a b‐roll with two strands per coil—has been proposed for HET‐s amyloid filaments (Ritter et al., 2005). (C) A set of b‐helical filament structures that would satisfy the mass‐per‐unit‐length requirement of one subunit per 0.47 nm that has been determined for Ure2p and some Sup35p filaments. Here, again, different colors denote different subunits. Left—one protofilament, one coil per subunit, six subunits; middle—two protofilaments, two coils per subunit, three subunits shown per protofilament; right—three protofilaments, three coils per subunit, two subunits shown per protofilament. The dotted and dashed segments are N‐ and C‐terminal linkers, eventually connecting to a globular domain (or domains).

molecular axis. A b‐helix, therefore, is a cross‐b structure. Most examples to date are monomeric proteins, but trimers of two kinds have been observed: threefold bundles of single‐stranded b‐helices (Steinbacher et al., 1994) and triple‐stranded b‐helices (Makhov et al., 1993; van Raaij et al., 1999). It has been suggested that the Sup35p filament may be a bundle of four cylindrical b‐sheets or nanotubes (Kishimoto et al., 2004). The nanotube is a hypothetical structure (Perutz et al., 2002) whose winding of the polypeptide chain is topologically similar to that of a b‐helix but it is round in cross section and water filled whereas b‐helices have cross sections with triangular or other shapes and water is largely excluded from their interiors (Kajava and Steven, 2006). The model of Kishimoto et al. envisaged six coils


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per subunit, and therefore, 0.67 subunits per 0.47‐nm axial step; in contrast, the observed values of 1.00–1.15 subunits per step (Diaz‐Avalos et al., 2005) are >50% higher, representing an unacceptably high discrepancy. Another proposal has been made that Sup35p filaments are b‐helical (Krishnan and Lindquist, 2005) but the model suggested by these investigators is nonpolar and therefore appears to be ruled out by the polarity data (Section V.A). Their experiments with Cys mutants and fluorescence labels and chemical cross‐linking on Sup35pNM filaments suggest that the head and tail regions from residues 25 to 38 and from 91 to 106 in 1 molecule are in close proximity to the same regions of their neighbors while the central region (residues 39–90) does not form intermolecular contacts. Neither polar b‐helical models nor a superpleated b‐structure that includes the central region can readily explain this interpretation of these data. Nevertheless, a superpleated b‐structure formed by the head and tail regions with the central region forming a long loop protruding from the fibril, or a model in which the head and tail regions each form coiled stacks with a single coil of b‐helix or b‐roll per subunit could be squared with this interpretation. We now consider some other formulations for b‐helical or b‐helix‐like (i.e., b‐roll or nanotube) models, representing generalizations of the model of Kishimoto et al. (2004) to different numbers of coils per subunit and protofilaments. Since there are typically 19–27 residues per coil of a b‐helix, the N‐domain of Sup35p at 114 residues could fold into, at most, 6 coils. However, the truncated N‐domain (residues 1–61) in the construct used by Diaz‐Avalos et al. (2005) suffices for a maximum of three coils. If we interpret their mass‐per‐unit‐length data as indicative of one subunit per 0.47‐nm axial step, then a filament could, in principle, have one coil per subunit, with axial stacking of the coils in successive subunits; or two protofilaments, each having two coils per subunit; or three protofilaments, each having three coils per subunit (Fig. 11). These models invoke axial stacking of b‐helical coils from different subunits within a protofilament, an arrangement that is theoretically possible but has not been demonstrated. The trimeric bundle has the advantage that there are precedents for such a structure, for example, the tailspike of phage P22 (Steinbacher et al., 1994). However, the disadvantage of this model, and indeed of all multicoil‐per‐subunit models, is that it does not invoke in‐register stacking of adjacent polypeptide chains. Accordingly, such arrangements are unlikely to survive sequence randomization (see above). The only such model to give in‐register stacking of adjacent chains has one coil per subunit (Fig. 11). However, it corresponds to a rather small portion of the prion domain forming amyloid—say, 25 residues—which would correspond to an amyloid core of 2.5 nm in diameter. This appears to be too narrow

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(cf. 5.0 0.9 nm as measured by us from Fig. 2 of Kishimoto et al., 2004). Moreover, the conformational change on amyloid formation would be very small (25 residues out of 685, i.e., 3.6%). Based on these considerations, we conclude that while current data do not entirely rule out models of this kind for Sup35p, they render them unlikely. In principle, models of this kind are also candidates for Ure2p filaments. In this case also, the observed preservation of the ability to fibrillize despite sequence randomization of the prion domain suggests that the one coil‐per‐subunit version is the only viable candidate. However, the protease‐protected region should be no more than about 25 residues which is inconsistent with the experimentally observed value of 65–70 residues (Baxa et al., 2003). Moreover, the b‐sheet content of Ure2p1–65 fibrils would be no more than about 40% which seems too low, given the experimental estimate of >65%, even with a large margin of error (Section IV.G and Table III). Accordingly, we consider such models unlikely. For HET‐s filaments, a b‐roll model has been suggested (Ritter et al., 2005) based on a secondary structure assignment by solid‐state NMR and hydrogen exchange studies that specified the positions of four b‐strands in the prion domain (Fig. 11). The authors detected sequence similarities between strands 1 and 3 and between strands 2 and 4. Based on that observation, they proposed pseudo in‐register interactions between respective strands in a structure that forms a b‐roll (a variant of the b‐helix with two strands per turn instead of three) with two coils per subunit and two b‐strands per coil. This model predicts an axial repeat of 0.94 nm (2 0.47 nm) and does not have in‐register packing of successive molecules assuming all molecules have the same conformation, that is, strand 1 interacts with strand 3 intra‐ and intermolecularly. The predicted mass‐ per‐unit‐length and the molecular interactions are properties that are measurable.

VII.

Other Structural Considerations

A. Crystal Structure of a Seven‐Residue Peptide from Sup35p Recently, a seven‐residue peptide from the Sup35p prion domain has been analyzed by X‐ray crystallography (Nelson et al., 2005). Although there is, in general, little reason to suppose that a short peptide will assume the same structure in a crystal as it will in the context of a folded protein containing it, these crystals seem to be related to amyloid fibrils of the same


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peptide (Diaz‐Avalos et al., 2003). In the crystal, the peptide assumes an extended conformation and is packed in a parallel in‐register arrangement that optimizes the possibility for stacking of Asn and Gln whereby their side chains can form H‐bonds. Pertinent observations from the crystal structure are as follows: (1) The peptides are packed so as to generate an array of parallel b‐sheets with in‐register stacking of the peptides as individual b‐strands. (2) There is a so‐called ‘‘dry’’ interface between sheets in the crystal, that is, the interacting surfaces of two b‐sheets have only polar residues packing to form a solvent‐free core. All possible hydrogen partners are saturated in this ‘‘polar’’ core either by in‐register stacking of like residues such as Asn or Gln or by contacts with neighboring residues in the same or in adjacent subunits. This arrangement is somewhat unorthodox because it is usually assumed that at least some hydrophobic residues are needed to form a compact solvent‐free core. (3) The b‐strands in adjacent parallel b‐sheets are oppositely oriented. (4) There is a 0.24 nm offset along the stacking axis between neighboring b‐sheets so that their strands are relatively staggered by one half of the 0.47‐nm increment. This arrangement optimizes the contact surface between the sheets. Comparison of this crystal structure with previously suggested models (Section VI) shows much in common with the superpleated b‐structure. In both the model and the crystal, there is in‐register packing of b‐strands within parallel b‐sheets, opposite orientations of the strands in adjacent b‐sheets, and location of polar residues (Asn and Gln) in a compact solvent‐free core, where these residues form a network of H‐bonds. In the model, neighboring sheets are in axial register whereas in the crystal structure, they are offset by 0.24 nm. It remains to be seen whether this discrepancy is real or whether it arises from constraints imposed in accommodating turns in a prion domain filament, but not in a peptide crystal. The strands of the b‐roll model for HET‐s filaments (Ritter et al., 2005) also have an arrangement that resembles the one in the crystal structure (parallel in‐register packing within b‐sheets; opposite orientations in adjacent sheets).

B.

Polymorphism in Prion Filaments

In many amyloid systems, filament polymorphism has been observed by EM (e.g., Goldsbury et al., 1997, 2000). Structural variations may be expressed in terms of long‐range axial repeats (Goldsbury et al., 2005; Jimenez et al., 2001), diameter (Louis et al., 2005), and/or number of protofilaments ( Jimenez et al., 2002). Solid‐state NMR has also been used to detect slight structural differences in Alzheimer’s b‐peptide filaments

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made under different physical conditions (‘‘quiescent’’ vs ‘‘agitated’’) (Petkova et al., 2005). Among fungal prions, filament polymorphism has been observed for Ure2p‐related constructs (Baxa et al., 2002) in the value of their long‐range axial repeat and the number of protofilaments (one or two). Polymorphism in the Sup35p system has been inferred from observations of seeding capacity and filament growth rate (DePace and Weissman, 2002; Uptain et al., 2001) and also on overall filament morphology and mass‐per‐unit‐length data (Diaz‐Avalos et al., 2005). The Ure2p N‐domain/GFP fusion turned out to be an especially fruitful construct, forming filaments with conspicuous axial repeats that vary from filament to filament (Baxa et al., 2002). We have investigated how this polymorphism may be affected by the length of the linker connecting the GFP appendage to the core fibril by comparing the following constructs: Ure2p1–65‐GFP, Ure2p1–85‐GFP, Ure2p1–90‐GFP, and Ure2p1–95‐GFP. Micrographs and histograms of observed repeat lengths are presented in Figs. 12 and 13. (1) For all constructs, when the repeat is clearly exhibited, its value is rather constant within a given filament but varies markedly between filaments (Fig. 12). (2) For all constructs we observe three major classes with short (60–70 nm), medium (90–115 nm), and long (>130 nm) repeats (except Ure2p1–65‐GFP, for which we do not find the long class). (3) The most common repeats are around 100 nm, although the peak of the distribution increases slightly with increasing linker length (80 nm for Ure2p1–65‐GFP, 95 nm for Ure2p1–85‐GFP, 97 nm for Ure2p1–90‐GFP, and 105 nm for Ure2p1–95‐GFP). These results indicate that the repeat length (and the underlying twist at each 0.47‐nm increment) is an intrinsic property of a given filament that is maintained as it grows longer. Importantly, they also document that in vitro ‐assembled filaments exhibit a range of structures, each probably based on a particular kind of nucleation event. This heterogeneity in filament structure correlates with the fact that the filaments give rise to a variety of variants in vivo when transformed into yeast cells (Section VIII.C). We infer that the spread of repeat lengths appears to be determined primarily by the region from residue 1 to 65. Although the linker between that region and the appended GFP moiety has some influence, it is not the principal structural determinant. With an axial rise per subunit of 0.47 nm (see above), these repeats translate into twist angles per subunit: for short repeats of about 40 nm, 4–5 per subunit; for the longest repeats of about 150 nm, 1.0 per subunit. The commonest repeat of 100 nm corresponds to a twist angle of 1.8 per subunit.


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Fig. 12. Polymorphism of filaments formed from the fusion protein constructs Ure2p1–65‐GFP, Ure2p1–85‐GFP, Ure2p1–90‐GFP, and Ure2p1–95‐GFP is illustrated with representative negatively stained micrographs of filaments. The panels are labeled (in white), according to the construct from which each filament was made. In projection, the filaments have a sinusoidal form whose wavelength is constant within a given filament but varies from filament to filament. The filament shown at top left is a double filament, consisting of two single filaments wrapped around each other. Bar ¼ 200 nm.

Polymorphism also seems to influence the growth rate of filaments. In their AFM study of Sup35pNM filaments involving a compilation of the growth rates measured on many individual filaments, DePace and Weissmann (2002) found different classes, for example, fast on both ends, slow at one end, and so on (Section V.A; Fig. 8). They were able to show in several rounds of growth analysis that growth rate is an intrinsic property for each class of filaments.

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Fig. 13. Distributions of repeat lengths in filaments of Ure2p‐GFP fusion constructs. Each count represents one filament, no matter how long. The standard deviation of repeats along a single filament was about 5 nm, on average and is shown inset into the Ure2p1–85‐GFP graph.

VIII.

Prion Variants

Strains or variants of TSEs are a well‐established phenomenon, with different isolates showing variations in incubation times, disease symptoms, and the brain regions affected in genetically identical hosts (reviewed by Bruce, 2003). This variability was used as an argument against the protein‐ only hypothesis (Carp et al., 1994) because it was not evident how the same protein could have multiple self‐replicating conformations. In the fungal systems, a similar phenomenon was first observed for [PSI] by Derkatch et al. (1996). To avoid confusion with strains of host organisms (yeast strains) and prion strains, we will use the term variants for phenotypically distinct versions of the same prion. Since the protein‐only hypothesis has been validated for fungal prions (Section III.E), it is now clear that these prion proteins have not only one infectious conformation but several—one for each variant. This important and intriguing feature of prions has to


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be accounted for by tenable structural models. Prion variants and their relationship with species barriers have been discussed in detail by Chien et al. (2004).

A. Detection Methods As mentioned above, TSE variants are distinguished by their incubation times and disease symptoms. For fungal systems, the only distinguishing biological properties to date are the residual activity of soluble protein (the ‘‘strength’’ of the prion, a strong variant having low residual activity) and the frequency of loss of the prion (its so‐called ‘‘mitotic stability’’). These two properties are somewhat related (Brachmann et al., 2005; Derkatch et al., 1996). For both properties, it is important for investigation to have a tunable selection system that not only selects for a given activity but also for the level of that activity. Another way to detect variants was developed by King (2001) who used point mutations in the prion domain of Sup35p to test their compatibility with each variant. Each variant has a specific pattern of compatibility with the point mutations used. The ADE marker is a good selection system for detecting variants used in the [PSI] system (Cox, 1965; Derkatch et al., 1996; Fig. 2) and, more recently, was adapted for [URE3] (Brachmann et al., 2005; Schlumpberger et al., 2001). In both cases, cells in the non‐prion state are Ade, lacking an enzyme in the adenine biosynthesis pathway. Cells in the prion state can be selected on medium without adenine by their Adeþ phenotype. However, when cells are grown on low‐adenine medium, all cells are able to survive and grow, but an intermediate of the adenine biosynthesis accumulates and is converted to a red pigment after spontaneous oxidation. That results in a strong red colony color in non‐prion cells and in a white color for prion‐containing cells. However phenotypes in between (different shades of pink) are possible and can be used to detect different variants in the prion systems (Brachmann et al., 2005; Derkatch et al., 1996; Schlumpberger et al., 2001).

B. Relationship Between Filament Polymorphism and Prion Variants A key feature of the protein‐only hypothesis is that variants should represent distinct structural forms of the prion protein. A direct connection between filament structure and variants was made by Tanaka et al. (2004) using filaments made under different conditions (in this case, 4 C and 37 C). Filaments formed at 4 C were less stable against heating in 1.6% SDS and gave rise mostly to strong [PSI] variants after being transformed

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Fig. 14. Prion variants may be propagated faithfully in vitro. (A) Scheme of propagation experiment. (B) Randomly selected transformants from experiments in which soluble Ure2p was seeded with extracts from [ure‐o] cells or [URE3] variant one, two, or three cells, respectively. Panel (B) was adapted from Fig. 5 of Brachmann et al. (2005).

into yeast, whereas filaments formed at room temperature or 37 C are more stable and give rise to weak [PSI] variants. This finding correlates well with the idea that phenotypically strong prion variants reflect physically less stable filaments that break more often, producing more growing ends. As a result, they are able to deplete the pool of soluble protein faster. King and Diaz‐Avalos (2004) found that solutions of soluble Sup35p1–61‐GFP seeded with extracts of yeast cells harboring a certain prion variant can transmit this variant faithfully through five passages of reseeding in vitro followed by transformation into yeast. A similar result was observed in the Ure2p system, but only one round of seeding was possible before the specific variant was lost (Brachmann et al., 2005; Fig. 14). These results clearly show that only the protein (specifically, the prion domain) is needed for variant‐specific infectivity. This method was used to produce homogeneous preparations of in vitro ‐assembled filaments of a single variant of Sup35p. These filaments have been studied by electron microscopy and some structural variability was detected (Diaz‐Avalos et al., 2005). In another study, distinct variants were inferred to have different structures by looking for seeding efficiency of yeast extracts (Uptain et al., 2001). Extracts from strong [PSI] variants were more efficient in seeding new filament growth than extracts from weak variants.


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C. In Vitro Filament Populations Are Heterogeneous Whereas Intracellular Filament Populations Are Homogeneous The possibility to transform in vitro ‐made filaments into yeast cells and observe different variants allows the homogeneity of these preparations to be assessed. If a preparation were to be homogeneous, transformation of it should result in only one variant. However, the experimental observations are to the contrary. Different shades from almost red through pink to completely white [URE3] variants were observed in transformed yeast clones from a single preparation (Fig. 6; Brachmann et al., 2005). Similar observations were made earlier with Sup35p filaments (Tanaka et al., 2004). These data imply that filament preparations made in vitro are mixtures of structures, each corresponding to a distinct variant. In vivo, however, spontaneous nucleation seems to be a very rare event, since variants are stable and are transmitted faithfully. Only very few observations of conversion from one variant to another have been reported. It seems that usually only one type of filament, resulting from a single nucleation event, resides in a yeast cell at a given time. Therefore, the filament population in a given prion‐infected cell is homogeneous. In the case that two variants are in a cell at the same time (e.g., after transformation with in vitro filaments), the stronger variant will be the only one that is detected by the assay and most likely also the only one that survives the competition since it grows faster and is transmitted more stable to the daughter cells.

D. How Are Variants Explained by the Models? All models that invoke parallel in‐register b‐sheet arrangements can adopt variations only in the topology of the peptide chain. Therefore, variants in these models should mainly be driven by side chain interactions. In the superpleated b‐sheet model, variations in the exact position of turns, turn conformations, strand lengths, and strand number might give rise to different variants (Fig. 10). Since most of the proposed H‐bonds involve intermolecular rather than intramolecular interactions, a subunit attaching to the growing end of the filament has to adopt the same configuration of strands and turns to get the most favorable interaction in the in‐register arrangement: that is, the end of the filament presents a flat template that guides the folding of the next subunit to be added, that subunit having been unfolded while in solution (Fig. 15). In this way, the specific structure of that variant is propagated along the filament as it grows. A different variant can only arise from a new and different nucleation process.

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Fig. 15. Templated assembly of amyloid filaments in two models. The scheme shows how two models of prion filaments can explain the faithful propagation of prion variants: (A) A super‐pleated b‐structure model. (B) A polymeric b‐helical model. In both cases, preexisting filaments (seeds) are shown in blue. Arrows indicate b‐strands and are connected by turns. In both scenarios, an additional subunit (shown in red), initially natively unfolded in solution, adds to the growing filament. In each case, the exposed end of the seed presents a two‐dimensional template that guides the folding of the newly recruited subunit according to an in‐register stacking arrangement. In this scheme, folding and assembly are tightly coupled. The newly added subunit assumes same conformation (from left to right) as subunits already present in the filament. Two structural variants of the super‐pleated b‐structure (for the same amino acid sequence) are shown in the top two rows (A), and two different variants of single‐coil b‐helices (one coil per subunit) are shown in the lower two rows (B).


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As for b‐helical models in the context of prion variants, we note that b‐helices encompass quite a wide variety of cross‐sectional shapes, each reflecting a different configuration of strands and turns in the coil (Hennetin et al., 2006; Kajava and Steven, 2006): thus it seems possible that different parts of the same prion domain could be assigned the role of coil‐former and they would have different coil geometries whose distinctions could give rise to variant prions. However, we expect that variability of the b‐helices with more than one coil per subunit—already unlikely on other grounds, except for HET‐s (Section VI.B)—will be quite limited. Finally, how would the phenotypes of variants, which represent different levels of partial activity, come about? In the Ure2p system, the C‐terminal functional domain is inactivated by being confined to filaments (Section IV.F). Irrespective of the precise structures of variant prion filaments, we can imagine two possible mechanisms: (i) different filament geometries are more or less efficient at recruiting from the soluble pool of Ure2p, leaving correspondingly sized pools of functional nonfilamentous protein, and (ii) the differing filament geometries allow, to varying degrees, the attached C‐terminal moieties to interact with the transcription factor Gln3p, and these varying degrees are reflected in characteristically different levels of partial activity.

IX. Perspective The fungal prion systems bring incisive practical advantages to the experimental investigation of prion phenomena, although they cannot be used to study the neurotoxicity of the PrPSc prion because they have no comparable effect on their host cells. Fungal prion proteins also provide useful models for the study of amyloid structures and their formation. The first discovered and, to this point, most extensively studied systems, [URE3] and [PSI], have turned out to be closely similar in almost all respects: in the unusual amino acid compositions and other properties of the N‐terminal prion domains of Ure2p and Sup35p; in the correlation between filament formation and prionogenesis; in the accompanying transition of the prion domains from unfolded to amyloid; and, in our view, probably also in the kind of amyloid structure that is formed in the filament backbone. Importantly, they are both loss‐of‐function prions. A third yeast prion, [PIN], is similar in some respects, but its non‐prion domain—in this case, the N‐terminal part of the molecule—has no established cellular function. The protein Rnq1p is a gain‐of‐function prion whose activity is expressed via its ability to induce [PSI]. [PIN] forms much more readily than the other two prions, and [PIN] cells induce [PSI] under

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conditions in which spontaneous generation of [PSI] cannot occur. In terms of the scenario outlined above, we conjecture that the prion domain of Rnq1p which is much larger than that of Sup35p but contains similar Q‐rich tracts may nucleate amyloid. These filaments may be able to nucleate extension with Sup35p from the template of those tracts. The [Het‐s] system of P. anserina has some basic differences from the other systems: (1) Its prion domain has a normal amino acid composition. (2) Although definitive structural solutions for any of the prion domain filaments have yet to be achieved, the body of experimental data is growing and it is possible that the fold and packing of prion domains in HET‐s filaments differ qualitatively from those in Ure2p or Sup35p filaments (Section VI). (3) Consistent with this proposition, variants have been reported for all the other systems but not for [Het‐s]. (4) [Het‐s] is a gain‐of‐function prion and the only prion currently on record to have an evolved cellular function, specifically, a key role in heterokaryon incompatibility. How this function is accomplished is unclear but it seems possible that the prion filaments may constitute some kind of platform on which allelic comparison can be performed.

Acknowledgments We thank Dennis Winkler, Naiqian Cheng, Joe Wall, and Martha Simon for their expert help and advice in various electron microscopy experiments; Anindito Sen and Sven Saupe for allowing us to cite unpublished work; Jonathan Weissman and Sarah Perrett for providing images; and Andreas Brachmann for comments on this chapter. This work was supported by the Intramural Research Program of the National Institute for Arthritis, Musculoskeletal and Skin Diseases and by a fellowship for T.C. from the Howard Hughes Medical Institute—National Institutes of Health Research Scholars Program.

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X‐RAY FIBER AND POWDER DIFFRACTION OF PRP PRION PEPTIDES By HIDEYO INOUYE AND DANIEL A. KIRSCHNER Department of Biology, Boston College, Chestnut Hill, Massachusetts 02467

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Prion Hypothesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Proteinaceous Infectious Particle Without Genome Is Causative Agent . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Structural Difference Between PrPC and PrPSc Arises in 90–145 Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Sequence Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Prion Alanine‐Rich Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Quarter‐Staggered b‐Chain Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. H1 Domain Forms a Reverse Turn. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Palindrome A8A Forms Intermolecular Hydrogen Bonding . . . . . . . . . D. 3F4 Epitope Concealed in PrPSc Exposed at Heterodimer Interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. His111 and Ala117 Interaction May Enhance Reverse ‐Turn Conformation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Cytotoxicity May Arise from Turn Conformation . . . . . . . . . . . . . . . . . . . . V. Amyloidogenic Core Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Polyalanine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Polyalanine Forms Cross ‐b Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Polyalanine Extension in Oculopharyngeal Muscular Dystrophy . . . . . VII. Polyglutamine. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Polyglutamine Forms b‐Crystallites But Not a b‐Helical Nanotube . . . B. Reverse Turn May Be Crucial for Fibril Formation . . . . . . . . . . . . . . . . . . VIII. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract A conformational change from the a‐helical, cellular form of prion to the b‐sheet, scrapie (infectious) form is the central event for prion replication. The folding mechanism underlying this conformational change has not yet been deciphered. Here, we review prion pathology and summarize X‐ray fiber and powder diffraction studies on the N‐terminal fragments of prion protein and on short sequences that initiate the b‐assembly for various fibrils, including poly(L‐alanine) and poly(L‐glutamine). We discuss how the quarter‐staggered b‐sheet assembly (like in polyalanine) and polar‐zipper b‐sheet formation (like in polyglutamine) may be involved in the formation of the scrapie form of prion. ADVANCES IN PROTEIN CHEMISTRY, Vol. 73 DOI: 10.1016/S0065-3233(06)73006-6

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I. Introduction Structural analysis of the prion protein includes the methods of solution NMR, X‐ray crystallography, electron microscopy, and fiber and powder diffraction. NMR solution studies of prions from different species (including human, hamster, bovine, cat, dog, pig, sheep, elk, chicken, turtle, and frog) show that the N‐terminal domain of the protein is flexibly disordered while its C‐terminal domain is an a‐helical globular structure (Calzolai et al., 2005; Donne et al., 1997; Gossert et al., 2005; Hornemann et al., 2004; James et al., 1997; Liu et al., 1999; Lopez et al., 2000; Luhrs et al., 2003; Lysek et al., 2005; Riek et al., 1996, 1997; Zahn et al., 2000). X‐ray crystallography also shows a globular C‐terminal domain (Haire et al., 2004; Knaus et al., 2001). Because spectroscopic data indicate that the cellular form of prion (PrPC) is largely a‐helical (Pan et al., 1993), the atomic structure of prion as determined by solution NMR and X‐ray crystallography is likely for the PrPC form. By contrast with these findings, a b‐sheet structure for prion (rods) and for the N‐terminal domain has been determined using X‐ray fiber and powder diffraction (Inouye and Kirschner, 1998; Inouye et al., 2000; Nguyen et al., 1995). Based on electron microscopy and spectroscopy, two different models have been proposed for the assembly of b‐chains in prion aggregates: (i) a parallel b‐helix (Govaerts et al., 2004; Wille et al., 2002) similar to that proposed for polyglutamine assembly (Inouye and Kirschner, 2005; Perutz et al., 2002a,b) and (ii) antiparallel b‐chains (DeMarco and Daggett, 2004; Sokolowski et al., 2003) or a polylysine‐like b‐structure (McColl et al., 2003; Padden et al., 1969). Conformational change from the cellular to the scrapie form is, perhaps, the central event in prion replication. A cell‐free study shows that the abnormal infectious form (PrPSc) which is predominantly in a b‐sheet conformation must be partially denatured for replication (Kocisko et al., 1994; Fig. 1). The denaturation exposes, in a strain‐specific manner, a particular sequence—residues MKHM corresponding to amino acids 109–112 in the Syrian hamster (or human) sequence—which is referred to as the 3F4 epitope (Safar et al., 1998; Fig. 2). This domain is likely to be localized at the binding interface between the cellular (PrPC) and scrapie forms (PrPSc). The known cytotoxicity of the PrP106–126 domain (De Gioia et al., 1994; Forloni et al., 1993) is likely structure dependent (Bergstrom et al., 2005). The binding of copper and zinc to His111 (Jackson et al., 2001) modulates prion’s aggregation and neurotoxic properties (Jobling et al., 2001). In particular, the hydrophobic palindrome AGAAAAGA (PrP113–120) has been shown to be necessary for prion propagation (Norstrom and Mastrianni, 2005). The sequence of residues in the PrP106–126 domain among many different animals is mostly

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v

Strain

v

v

v

3F4 epitope exposed

GdnHCI

GdnHCI

3F4 epitope exposed

Species barrier

In vitro replication PrPSc + [35S]PrPC 3 M GdnHCI

[35S]PrPSc 0.75 M GdnHCI

Fig. 1. Transmission mechanisms. Strain barrier: PrPC (circle) interacts with different strains of PrPSc (square or triangle). The replicated PrPSc is similar to the template. The 3F4 epitope is not recognized when it is in PrPSc, but is exposed after partial denaturation by GdnHCl so that it is detected by the antibody. Antibody reactivity depends on the particular strain of PrPSc (Safar et al., 1998). Species barrier: when the template PrPSc contains unfavorable residues at the binding interface, the transformation of PrPC to PrPSc does not occur. In vitro replication: 35S label of PrPC is detected in PrPSc after replication in a medium containing GdnHCl (Kocisko et al., 1994).

homologous; however, in the cervids, which include elk and deer, there is a Met112Val substitution in the 3F4 epitope domain (Raymond et al., 2000). This difference may hinder the heterodimerization of PrPcwd (the infectious prion in chronic wasting disease) with human prion, and consequently make PrPcwd‐induced conversion of human prion in a cell‐free system less efficient than that of elk or deer (Raymond et al., 2000). To address the issues of pathogenesis, prevention, transmission, and inactivation of prion diseases, it is crucial to understand the structure and conformational change involving the binding interface—that is, sequence 106–126 in SHa (or human), the homologous sequence (residues 109–129) in cervids, and analogues of these sequences. Fibrillar forms of biological macromolecules—such as collagen, DNA, muscle proteins, and filamentous viruses—have been elucidated using fiber and powder diffraction methods (Fraser and MacRae, 1973).

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1 11 21 31 41 51 MANLSYWLLA LFVAMWTDVG LCKKRPKPGG WNTGGSRYPG QGSPGGNRYP PQGGGTWGQP Signal * * 61 71 81 91 HGGGWGQPHG GGWGQPHGGG WGQPHGGGWG QGGGTHNQWN

* 121 VVGGLGGYML

181 NITIKQHTVT CHO H3 241 VILLISFLIF

*

*

A8A 101 111 KPSKPKTNMK HMAGAAAAGA H1

131 141 151 161 171 GSAMSRPMMH FGNDWEDRYY RENMNRYPNQ VYYRPVDQYN NQNNFVHDCV H2 S1 A S2 B 191 TTTKGENFTE

CHO 251 LMVG

201 TDIKIMERVV

211 221 231 EQMCTTQYQK ESQAYYDGRR SSAVLFSSPP H4 Gpi C

Primary structure of hamster PrP

Fig. 2. Primary structure of hamster PrP (Stahl et al., 1993). The first 22 residues at the N‐terminus are the signal sequence. PrPC is completely digested by proteinase K, whereas the N‐terminal sequence of PrPSc to residue 89 (arrow, closed head) is digested. CHO indicates the glycosylation sites at residues 181 and 197; Gpi the glycosylphosphatidylinositol anchor at 231; and * the N‐terminal octarepeats. In one case of human prion disease, a stop codon was found at 145 (arrow, open head) (Kitamoto et al., 1993). H1, H2, H3, and H4 denote the predicted a‐helices (Huang et al., 1994, 1996), and A–C denote the a‐helices and S1, S2 the b‐strands determined by solution NMR (James et al., 1997).

Fiber X‐ray patterns have also been obtained from a variety of amyloidogenic polypeptides including b‐amyloid, prions, polyalanine, and polyglutamine. Because these fibrillogenic materials are of defined composition, it is possible to build physically plausible molecular models of them from the dimensions revealed by their diffraction patterns. For fiber diffraction, the fiber axis is a cylindrical axis, about which the molecular assemblies have different rotational orientations. In a fibril having a cross‐b structure, the cylindrical axis is in the hydrogen‐bonding direction of the b‐sheet and is the direction of the fibril. The coherent length or domain size in a fibril— that is, the number of the lattice points—is largest in the fiber direction. Sheet‐like or slab‐like b‐structures can also give fiber patterns. When a sheet is extended in the hydrogen‐bonding and intersheet directions, the cylindrical axis or fibril axis is the direction of the b‐strands. The fiber axis does not always give the largest coherent length. Thus, for a slab‐like structure the cylindrical axis (the chain direction) corresponds to the smallest coherent length. Both fibril and slab‐like structures are observed for Alzheimer’s b‐amyloid analogues (Inouye et al., 1993) and for prion peptides (Inouye and Kirschner, 1997; Inouye et al., 2000; Nguyen et al., 1995). Fiber and


185

powder diffraction patterns often show lattice disorder and diffuse scattering arising from substitution disorder. Analyses of these have been presented previously (Guinier, 1963; Hosemann and Bagchi, 1962; Vainshtein, 1966; Worthington and Inouye, 1985), and references to disorders in fibrils are described in Inouye (1994). In this chapter, we review current knowledge of prion pathology (Figs. 1 and 3) and summarize the results of X‐ray fiber and powder diffraction applied to the alanine‐rich domain of prion, including the sequences SHa106–122 (KTNMKHMAGAAAAGAVV), H1 (SHa109–122), and A8A (SHa113–120) (Inouye and Kirschner, 1997, 1998; Inouye et al., 2000; Nguyen et al., 1995). To explore the common fibril formation mechanism involving b‐crystallites,

Fig. 3. Classification of human prion diseases. Sporadic: the transformation from PrPC (circle) to PrPSc (square) occurs without apparent cause. Familial: a point mutation (*) is thought to facilitate the transformation. Infectious: the transformation arises via PrPSc which acts as a template. The kinetic equations are defined by Eigen (1996). The infectious form includes kuru, iatrogenic CJD (iCJD), variant CJD (vCJD; first reported in 1996), bovine spongiform encephalopathy (BSE; first reported in 1985), and scrapie. In the nucleation‐dependent model, monomeric PrPC and PrPSc are in chemical equilibrium.

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we also compare structures of prion peptide assemblies with those of polyalanine and polyglutamine.

II. A.

Prion Hypothesis

Proteinaceous Infectious Particle Without Genome Is Causative Agent

The human prion diseases are classified as sporadic, familial, and infectious diseases (Fig. 3). The molecular mechanism in all cases underlying the protein‐only etiologies is the structural alteration from a‐helical cellular form of prion (PrPC or protease‐sensitive PrPsen) to the b‐sheet abnormal, infectious form (PrPSc or protease‐resistant PrPres) (Cohen et al., 1994; Horwich and Weissman, 1997; Prusiner, 1982, Prusiner et al., 1983). The sporadic type refers to the spontaneous structural transformation of PrPC ! PrPSc. The familial type indicates that a certain mutation may facilitate this PrPC ! PrPSc transformation. This includes Gerstmann‐ Straüssler‐Scheinker (GSS; with amino acid changes P102L, P105L, A117V, Y145Stop, F198S, Q217R), familial CJD (V180I, E200K, M232R) (Inouye et al., 2000; Salmona et al., 2003), and fatal familial insomnia (FFI) (D178N). The infectious type includes kuru, iatrogenic CJD, variant CJD (vCJD) for human (Bruce et al., 1997; Hill et al., 1997; Will et al., 1996), bovine spongiform encephalopathy (BSE) for bovine, and scrapie for sheep (Prusiner, 1982, 1991). There are many incidents of chronic wasting disease with amyloid fibrils for mule deer (Guiroy et al., 1993a; Williams and Young, 1980) and elk (Guiroy et al., 1993b; Williams and Young, 1982). Consistent with the protein‐only scenario of prion disease etiology, aggregated synthetic prion peptide has now been shown to be infectious (Legname et al., 2004). In the same manner that flagella from different bacterial strains give different fibril morphologies (Oosawa and Asakura, 1975), prion fibrils also have varied morphologies, as shown by atomic force microscopy (ATM) and spectroscopy (Jones and Surewicz, 2005). Stain‐specific morphologies of yeast prion amyloid fibrils are also indicated from mass per length measurements (Diaz‐Avalos et al., 2005). The fact that prion disease can be infectious indicates that the endogenous PrPC must be transformed by exogenous PrPSc which acts as a template. This prediction is confirmed by the observation that no infection is found in transgenic mice that lack the endogenous prion gene (Bueler et al., 1993; Prusiner et al., 1993). Two mechanistic models of PrPSc amplification have been proposed—the heterodimer model (Cohen et al., 1994) and the nucleation‐dependent model (Jarrett and Lansbury, 1993) (Fig. 3).


187

In the former, normal and abnormal prion molecules interact with each other, while in the latter, normal and abnormal prions are in chemical equilibrium, and the abnormal prions can form an oligomer. Also, in the nucleation‐dependent model the PrPSc is simply an aggregated form of PrPC, where the rate of oligomerization depends on the initial oligomer size. If the oligomer is smaller than the initial nucleus, monomer addition to oligomer is slow, but if the oligomer is larger than the nucleus then the reaction proceeds rapidly. This model, therefore, predicts that if the exogenous PrPSc aggregate is larger than that of the nucleus, then PrPC molecules are integrated rapidly into a PrPSc fibril. Equations and kinetic constants for quantifying the number of monomers in the nucleus and fibril have been derived (Eigen, 1996; Ferrone, 1999; Oosawa and Asakura, 1975) and these have been related to the pH‐dependent binding of Congo red to amyloid (Inouye and Kirschner, 2000). The nucleation‐dependent model, which posits that the infectious PrPSc particle is simply a polymeric (fibrillar) form of PrPC, is at variance with the observations that prion infectivity arises from a small oligomer (Silveira et al., 2005) and is unrelated to fibril formation (Wille et al., 1996, 2000).

B.

Structural Difference Between PrPC and PrPSc Arises in 90–145 Domain

The primary structures of PrPC and PrPSc are identical (Fig. 2), with both—for example, hamster prion precursor sequence SHaPrP (Stahl et al., 1993)—composed of 254 amino acids. The first 22 residues at the N‐terminus comprise the signal peptide. The prion protein is attached to the cell membrane at Ser231 via glycosylphosphatidylinositol (GPI). After the N‐ and C‐termini are cleaved, the prion consists of the sequence from residues 23 to 231. The repetition of hexameric homologous sequences within the N‐terminal sequence 51–90 provides binding sites for copper (Burns et al., 2002; Jackson et al., 2001; Viles et al., 1999). Secondary structure prediction suggests that there are four a‐helical domains (denoted as H1, H2, H3, and H4; Nguyen et al., 1995). At H1’s N‐terminus, the sequence MKHM (SHa109–112) has been identified as the epitope for the 3F4 antibody (Peretz et al., 1997) that can inhibit prion propagation (Peretz et al., 2001). This antibody has also been used to identify PrPSc immunochemically in tissue (Giaccone et al., 2000). H1’s C‐terminal half is rich in alanine (i.e., AGAAAAGA) and is denoted by A8A. The structural difference between PrPC and PrPSc is highlighted by their different sensitivities to digestion with proteinase K, whereas PrPC is digested completely, PrPSc generates the undigested 27‐ to 30‐kDa domain consisting of residues 90–231.

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This domain, denoted by PrP27–30, retains infectivity and forms amyloid fibrils (or prion rods) after detergent treatment (Prusiner et al., 1983). Spectroscopy, including CD and FTIR, shows that the secondary structure of PrPC is mostly a‐helical while that of PrPSc is b‐sheet (Pan et al., 1993). What specific regions within the whole of PrP are folded differently between PrPC and PrPSc? On the one hand, sequence 90–145 (generated in a GSS patient where a point mutation yields Y145Stop) forms b‐rich amyloid deposits (Kitamoto et al., 1993). Further, epitope mapping pinpoints a structural difference within the sequence 90–120 (Peretz et al., 1997). By contrast, NMR spectroscopy shows that the 90–145 domain within SHa90–231 is flexibly disordered (James et al., 1997) (Fig. 4), and presumably able, therefore, to assume either an a‐helical or b‐conformation depending on environment. Thus, the N‐terminal domain is likely to be where structural conformation differs between PrPC and PrPSc.

Fig. 4. The molecular structure, determined by solution NMR (James et al., 1997), of Syrian hamster 90–231 (SHa90–231) prion with ball‐and‐stick representation of the H1 domain (SHa109–122; MKHMAGAAAAGAVV). Note that two short b‐chains (S1, S2) nearly stack in the hydrogen‐bonding direction. If the palindromic polyalanine region was also in a b‐conformation, there would be a three‐stranded b‐sheet. The structural difference between PrPC and PrPSc is in the 90–145 domain. [Model drawn using MOLSCRIPT (Kraulis, 1991)].

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III.

Sequence Analysis

Pairwise comparison of PrP sequences among different animals (including human, bovine, sheep, mule deer, elk, golden hamster, and mouse) (Tables I and II) indicates that the human sequence in the core region (i.e., residues 90–230) is similar to that of other animals in the order: human > bovine > sheep > mule deer ¼ elk > mouse > golden hamster. At the putative heterodimerization interface, human106–126 is homologous to golden hamster or Syrian hamster SHa106–126; and the Met112 (human) ! Val substitution is present in the sequences of bovine, sheep, mule deer, and elk. The Met109 and Met112 in human prion are substituted in mouse by Leu and Val, respectively. The substitution Met ! Val in cervids at the putative‐binding interface likely accounts for the finding that the PrPcwd‐induced conversion in a cell‐free system is less efficient for human prion than for deer or elk (Raymond et al., 2000). Physical–chemical characteristics of prion proteins are evident after assigning different parameters, including hydrophobicity, charge, and molecular weight to the amino acid residues (Inouye and Kirschner, 1991, 1998) (Table II, Fig. 5). The sequences among different animals

Table I Pairwise Alignment of Prion Sequences (Identity Between Species in %)a

Human Bovine Sheep Mule deer Elk Golden hamster Mouse

Human

Bovine

Sheep

–

88.6 (92.2) –

89.9 (90.8) 94.3 (95.0) –

Mule deer 89.5 (90.1) 94.3 (95.0) 97.8 (95.7) –

Elk 89.5 (90.1) 94.3 (95.0) 97.7 (95.7) 99.2 (98.6) –

Golden hamster 89.4 (88.9) 84.9 (87.3) 87.2 (88.0) 87.2 (88.0) 87.2 (88.0) –

Mouse 89.0 (88.8) 84.6 (88.1) 86.5 (88.1) 86.5 (88.1) 86.5 (88.1) 93.7 (93.7) –

a Comparison by pairwise alignment (http://molbiol.soton.ac.uk/compute/align. html; Pearson and Lipman, 1988) between different species (including signal sequence). Comparisons of core sequences (residues 90–230) are shown in parentheses. The code numbers for the sequences, obtained from the Swiss protein databank, are: P04156 (human), P10279 (bovine), P23907 (sheep), P47852 (mule deer), P79142 (elk), P04273 (golden hamster), and P04925 (mouse).

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Table II Physical–Chemical Parameters for Human Core Domain PrP90–230 and Homologous Regions in Different Animalsa

N Sequence # MW pI Type

Human

Bovine

Sheep

Mule deer

Elk

Golden hamster

Mouse

141 90–230 16043 8.5 0.12 aþb

140 102–241 15875 8.9 0.11 aþb

140 94–233 15953 9.2 0.12 aþb

140 94–233 15998 9.1 0.12 aþb

140 94–233 15972 8.8 0.11 aþb

142 90–231 16225 9.1 0.15 aþb

143 89–231 16268 9.1 0.13 aþb

a

The pairwise alignment was performed as earlier (Table I). Alignment between human and bovine included gaps on the N‐terminal side, as the number of pentapeptide repeats in bovine is longer by one; therefore, N‐terminal sequences were aligned manually. The H and OH atoms are not included in the calculations. N, number of amino acids; Sequence #, numbered region in full‐length protein; MW, molecular mass in Da; pI, isoelectric point (calculated); , hydropathy score; Type, secondary structure classification.

Charge H1

H2

H3

H4

Hydrophobicity

a-Helix

b-Strand S1

a-Helical amphiphilicity

0

A S2

B

C

Coil

Turn

50

b-Strand amphiphilicity

100 150 200 Hamster PrP residue number

250

Fig. 5. Physical–chemical parameters as a function of residue number for hamster PrP (Inouye and Kirschner, 1998). The parameters (arbitrary scale) are: charge at pH 7; hydrophobicity; a‐helix (solid), b‐strand (dashed); turn (solid), coil (dashed); a‐helical (solid) and b‐strand amphiphilicity (dashed). The predicted helices (Huang et al., 1994) are labeled H1, H2, H3, and H4, and the NMR‐observed helices and b‐strands are A–C and S1, S2, respectively (James et al., 1997).


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indicate that the prion has an a þ b type conformation. The complete sequence can be divided into two parts: N‐terminal 90–145 and C‐terminal 145–230. The former has a very alkaline pI, while that of the latter is neutral. At physiological pH, therefore, the N‐terminal region has a large positive charge, which may account for its flexibly disordered structure by NMR, while the C‐terminal domain is globular (Fig. 4).

IV. Prion Alanine‐Rich Domain A.

Quarter‐Staggered b‐Chain Assembly

X‐ray diffraction patterns from the assemblies formed by several prion‐ related peptides have been analyzed (Fig. 6), including SHa106–122 (KTNMKHMAGAAAAGAVV) and H1 (i.e., SHa109–122, MKHMAGAAAAGAVV) (Inouye et al., 2000; Nguyen et al., 1995), and the palindromic A8A

(201) (002) (200)

*

(112)(211)

(201)

A8A (202)

(002)

Intensity

(004) (201) (200)

(004)(104)

H1(L) (302)

(002) (201)

(002) (200) (100)

* 0

*

H1(S/D) (300)

(204)

106−122 0.1

0.2

0.3

0.4

Reciprocal coordinate (1/Å)

Fig. 6. X‐ray diffraction intensity (arbitrary units) as a function of reciprocal coordinate for prion peptides. 106–122: meridional scan of the pattern of SHa106–122 dried from 50% AcN; H1(S/D): meridional scan of the pattern of SHa109–122 dried from 50% AcN; H1(L): lyophilized H1; A8A: SHa113–120 dried from 50% AcN. SHa106–122 is from Fig. 2A in Inouye et al. (2000). Diffraction patterns of A8A, H1(L), and H1(S/D) were previously reported (Nguyen et al., 1995). The strongest reflections (with Miller indices) are: 4.56 A˚ (201) in SHa106–122, 4.77 A˚ (200) in H1 (S/D), 4.44 A˚ (201) in H1 (L), and 4.33 A˚ (201) in A8A. Low‐angle reflections arising from the stacking of slabs are indicated by *.

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sequence (SHa113–120, AGAAAAGA). Oriented patterns were obtained from SHa106–122 and SHa109–122 dried from 50% acetonitrile (AcN) whereas powder patterns were obtained from lyophilized H1 and A8A dried from 50% AcN. The wide‐angle reflections correspond to a two‐dimensional, nearly orthogonal lattice having unit cell dimensions a ¼ 9.50–9.58 A˚ and c ¼ 10–16 A˚ (Table III). The a‐axis is in the direction of the hydrogen bonding for the b‐sheet, and c‐axis is in the intersheet direction. In the slightly oriented patterns from SHa106–122 and H1 assemblies, the c ¼ 14–16 A˚ which is larger than the 10–12 A˚ for the powder patterns of lyophilized H1 and A8A. The reflections from the two‐dimensional lattice are accentuated in the same direction, indicating that the cylindrical axis is along the b‐chain and the scattering object is slab‐like. The wide‐angle (201) reflection is strong, whereas the (200) reflection at 4.7 A˚ Bragg spacing is much weaker or nonexistent for A8A and lyophilized H1, or is nearly as strong as the (201) reflection for SHa106–122 and solubilized/dried H1. Because the intersheet distance of polyalanine is 5 A˚, the unit cell should contain two b‐sheets. The structure factor F(h,k,l) of Miller index hkl is given by X F ðh; k; lÞ ¼ fj exp½i2pðhx j þ ky j þ lz j Þ ð1Þ j

where fj is the atomic scattering factor at fractional coordinate (xj,yj,zj). The Miller indices hkl (integers) are defined in the hydrogen‐bonding, b‐chain, and intersheet directions. When the neighboring b‐sheets are quarter‐staggered in the hydrogen‐bonding direction, the fractional coordinates of the b‐chains are (0,0,0), (1/2, 0,0), (1/4,0,1/2), and (3/4,0,1/2). The structure factor is then written as 2 0 13 h l F ðh; 0; lÞ ¼ F0 ðh; 0; lÞ41 þ expi2p@ þ A5 4 2 2 0 13 3h l ð2Þ þG0 ðh; 0; lÞ4expiph þ expi2p@ þ A5 4 2 where F0 is the Fourier transform of one single b‐chain and G0 is the one for the antiparallel b‐chain. Using this equation, the structure factors can be calculated as F(2,0,0) ¼ 0 and F(2,0,1) ¼ 2[F0(2,0,1) þ G0(2,0,1)]. Thus, the absence of the 4.7 A˚ [indexed as (200)] and the strong (201) reflection suggests a quarter‐staggered b‐chain arrangement as proposed for b‐silk (Marsh et al., 1955a,b). The presence of the (200) intensity suggests that the b‐sheets are not exactly quarter‐staggered or that the structures of

Peptide Sequence Preparationa a (A˚) b (A˚) c (A˚) Angle Robs‐amp

H1 SHa109–122 L 9.58 – 11.84 90 0.36

H1 SHa109–122 AcN 9.51 7.06 15.94 b ¼ 88.4 0.24

A8A AGA4GA AcN 9.52 6.33 10.25 90 0.36

SHa106–122 SHa106–122 AcN 9.50 – 14.35 90 0.34

PolyA7 AcKYA7KNH2 S/D 9.61 6.91 11.33 90 0.25

PolyQ8 AcQ8NH2 L 9.73 7.14 8.16 g ¼ 95.7 0.24

a L, lyophilized; AcN, dried from 50% acetonitrile; S/D, solubilized in water, then dried. Dried H1 and lyophilized Q8 are monoclinic, while others are orthogonal. References for data: prion, Inouye and Kirschner (2003); polyA7, Shinchuk et al. (2005); polyQ8, Sharma et al. (2005).


Table III Summary of Crystal Data for Prion Peptides

193

194


Fig. 7. Ribbon models (upper) and electron density projections (lower) based on structural analysis of X‐ray fiber diffraction patterns of H1 (SHa109–122) (Inouye and Kirschner, 2003). (A) Antiparallel, (B) parallel, and (C) staggered arrangement of H1 dried from 50% AcN, and (D) lyophilized H1. Upper: the molecular model (Kraulis, 1991) which fits the observed electron density map shows two b‐chains, where Gly114– Ala115 are at the inverse turn (as defined by STRIDE; Frishman and Argos, 1995). Lys110, His111, and Ala117 are indicated by ball and stick models. The Lys side chains (and those of His) are close together in the antiparallel and parallel arrangements, whereas Lys and His are adjacent to one another in the staggered arrangement (C). In the lyophilized sample (D), the alanine‐rich domain modeled as a b‐strand fits the electron density distribution, and the 3F4 domain is likely disordered (not shown). Lower: XtalView representations of skeletal models superposed on the electron density projection along the chain direction of H1. H bonding is horizontal, and the intersheet direction is vertical.

four b‐chains in the unit cell are not identical. The intersheet distance is 8 A˚ for SHa106–122 and solubilized/dried H1, but 5 A˚ for lyophilized H1 and A8A. Using the b‐silk backbone as an initial phase model, the structure amplitudes from the model were calculated, and compared with the structure amplitudes extracted from the observed intensity. The electron density maps from the observed structure amplitudes and phases from the silk model for SHa106–122 and solubilized/dried H1 show electron density peaks other than those expected from the peptide backbone. For the lyophilized H1 and A8A, the b‐silk backbone fits the observed electron density map. Molecular models of prion peptides in which side chains are included fit the electron density maps even better (Fig. 7).

B.

H1 Domain Forms a Reverse Turn

In H1 the intrachain turn creates two antiparallel b‐strands, one consists of larger residues at the N‐terminus and the other is the polyalanine


195

region. This structure agrees with the slab thickness as measured from the low‐angle diffraction and the electron density peaks. Three molecular packings of H1 and SHa106–122 dried from AcN are proposed (Inouye and Kirschner, 2003; Inouye et al., 2000) (Fig. 7): (i) H1 molecules are in‐ register, and the b‐chains are arranged antiparallel; (ii) H1 molecules are in‐register, and the b‐chains are in a parallel arrangement; and (iii) H1 molecules are staggered in the intersheet direction and the b‐chains are antiparallel. The R‐factors (Ramp‐obs) are 0.46 for the antiparallel in‐register arrangement, 0.37 for parallel in‐register, and 0.24 for the staggered. A similar staggered assembly for the reverse turn in SHa106–122 gives an R‐factor ¼ 0.34. The atomic model of the polyalanine domain in lyophilized H1 (Fig. 7D) and in A8A give the same R‐factors ¼ 0.36. These calculations, which compare intensities calculated from the atomic models against the observed X‐ray intensities, suggest that the H1 domain likely forms a reverse turn, but under certain conditions the N‐terminal domain of H1 can become disordered such that b‐sheets are formed only by the C‐terminal alanine‐rich domain.

C.

Palindrome A8A Forms Intermolecular Hydrogen Bonding

In both the antiparallel and parallel b‐chain models, the histidine side chains are in close proximity to one another as are the lysine side chains (Fig. 7). Because of electrostatic repulsion between the lysine side chains, the in‐register arrangements may not be stabilized; rather, aromatic p–p interaction (Hunter and Sanders, 1990) between the imidazolium rings of histidine and polar‐zipper type side chain interaction between asparagine residues may stabilize this folding. In the staggered arrangement, the side chains of histidine and lysine residues are close together. If the positive‐ charged e‐amino of lysine is directed toward the aromatic ring of the histidine, the staggered arrangement may become stabilized by cation–p interaction as demonstrated in high‐resolution protein crystal structures (Burley and Petsko, 1986). In such an arrangement, His111 is closely positioned to Lys110 of the neighboring molecule. When a larger valine residue is introduced, for example in the GSS mutation Ala117Val, the His111 will interact with the valine more strongly than with lysine due to van der Waals contacts. Thus, in this GSS mutation the folded b‐sheet structure may be favored.

D. 3F4 Epitope Concealed in PrPSc Exposed at Heterodimer Interface To account for the species barrier of prion infectivity, it was proposed that the 109–112 epitope recognized by antibody 3F4 is localized at the

196


PrPC‐PrPSc binding interface (Scha¨tzl et al., 1995; Warwicker, 1997). The structural nature of the binding interface has been characterized by identifying peptides that inhibit prion replication in a cell‐free system. Such peptides presumably block the PrPC–PrPSc binding by interacting with the heterodimer interface. Prion‐related peptides SHa119–136, SHa166–179, and SHa200–223 inhibit the binding (Horiuchi et al., 2001) as does SHa106–141 (Chabry et al., 1998). That the 3F4 epitope in PrPSc is not immunoreactive (Peretz et al., 1997) suggests that the molecular conformation of PrPSc must be changed so as to expose this epitope when PrPSc interacts with PrPC. Based on the X‐ray fiber diffraction analyses, two different molecular structures for H1—folded in a b‐chain or disordered— may correspond, respectively, to the concealed and exposed 3F4 epitope.

E. His111 and Ala117 Interaction May Enhance Reverse ‐Turn Conformation To elucidate the roles of His111 and GSS mutation Ala117 ! Val in prion aggregation and toxicity, Salmona et al. (1999) studied SHa106–126 (KTNMKHMAGAAAAGAVVGGLG) analogues, including: (i) PrP106– 126HD, with l‐His111 ! d‐His substitution; (ii) PrP106–126A, with His111 ! Ala substitution; (iii) PrP106–126K, with His111 ! Lys substitution; (iv) PrP106–126V, with Ala117 ! Val substitution; (v) PrP106– 126NH2, with C‐terminal amidation; and (vi) PrP106–126VNH2, with Ala ! 117Val substitution and C‐terminal amidation. From spectroscopic and turbidity measurements of these peptides at pH 5.0 and 7.0, Salmona et al. (1999) suggest that His111 plays a central role in the conformational changes of PrP peptides. This agrees with the X‐ray model that His111 at neutral pH likely interacts with Ala117, while protonated His111 at acidic pH does not (Inouye and Kirschner, 1998). Although the X‐ray study suggests that the turn region contains residues Gly114–Ala115, spectroscopic data suggest that the turn is at Asn108‐Met112 (Ragg et al., 1999). The NMR data (James et al., 1997; Liu et al., 1999) show a slight reverse turn in the H1 domain, similar to that proposed from X‐ray diffraction (Inouye and Kirschner, 1998); however, NMR indicates that the turn is close to Ala117. A molecular dynamics study of the helix‐coil transition of PrP106–126 (Levy et al., 2001) indicates that the turn is near Ala115, such that His111 would interact with Val122 rather than with Ala117. The H1 domain, initially modeled as an a‐helix, also adopts a b‐hairpin fold as shown by molecular dynamics simulation (Daidone et al., 2005). Previous molecular modeling suggests that the heterodimer interface is at residues Ala118–Ala133 (Warwicker, 2000). As modeled, this domain is


197

an antiparallel b‐chain with a turn at Gly126–Gly127. Side chain interactions in the interesheet direction are via H bonding between the S atom of Met129 and the NH in Gly124, and van der Waals interactions between Val122 side chains. This model, however, agrees with neither the X‐ray model for H1 (109–122) (Inouye and Kirschner, 1998; Nguyen et al., 1995) nor the crystallographic results for PrP104–113 which shows a turn (Kanyo et al., 1999). Neighboring b‐strands in most proteins, including the PrP dimer proposed by Warwicker (2000), are hydrogen‐bonded together; however, there are examples in which neighboring b‐chains interact across the intersheet space—for example, in the H1 domain (Fig. 7), in the O‐turn of the Fab3F4 epitope in prion (Kanyo et al., 1999), in turn II (residues 7–20) of myelin P0‐glycoprotein (Shapiro et al., 1996), in Ab1–40 (Tycko, 2003), and in a model for Ure2p prion filaments (Baxa et al., 2005; Kajava et al., 2004). Molecular dynamics simulation of PrP106–126 also shows a reverse turn which enables the two b‐strands to interact with one another via their side chains across the intersheet space (Levy et al., 2001).

F. Cytotoxicity May Arise from Turn Conformation PrP106–126 is toxic to cultured neurons (Forloni et al., 1993) owing to its aggregation (Hope et al., 1996) or to its forming an ion channel (Kourie, 2001; Lin et al., 1997). Because this region of PrP contains the H1 domain (Fig. 2), it is likely that it too will form a b‐sheet structure. The sequences of the cytotoxic domains of Alzheimer’s Ab25–35 and amylin20–29 (Glenner et al., 1988) show sequence homology; and PrP106–126 shows some homology to these two: that is, the N‐terminal Ser(Thr)‐Asn and C‐terminal Ala‐Ile(Val)‐ Ile(Val).1 It is clear that the N‐terminal Asn may be structurally similar, in fact, to the ones found in the b‐chain ladder (Kobe and Deisenhofer, 1993; Yoder and Jurnak, 1995). Peptide

Sequence

Ab25–35 Amylin20–29 PrP106–126

GSN KGAIIGLM SNNFGAILSS KTNMKHMAGAAAAGAVVGGLG

Among different Ab analogues, full sequence Ab1–40 and fragment Ab25–35 are cytotoxic as fibers (El Khoury et al., 1996; Yan et al., 1996). 1 The homology between Ab25–35 and amylin20–29 was first recognized by D. A. Kane (unpublished observations).

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Because both present structurally as twisted fibers (Malinchik et al., 1998), it may be that this type of morphology has a high propensity for toxicity. As indicated by secondary structure predictions and solution NMR (Sticht et al., 1995), the hydrophilic part is largely turn or coil, and the hydrophobic part is an extended b‐chain or (in solution) a‐helical. The Ab22–35 hydrophilic turn (Tycko, 2003) might then be exposed on the surface of the Ab1–40 fiber. When Ab25–35 peptides are reconstituted with lipids, the peptide is localized within the hydrocarbon (Mason et al., 1996), which is consistent with the idea that the peptide assembly can act like an ion channel; however, this structure is not in agreement with hydrophilic localization of the turn. Powder X‐ray diffraction of Ab31– 35, which is homologous to tachykinin family of neuropeptides, shows that this region gives a reverse‐turn structure similar to one found for substance P (Bond et al., 2003). Tachykinins bind to G‐protein receptors, but it is not clear whether a similar receptor is present for binding PrP106–126. From the known correlation between the turn conformation of the hydrophilic residues of amyloidogenic peptides and their cytotoxicity, the cytotoxic effect of PrP106–126 likely arises from the turn region at 114–115 (Fig. 7).

V. Amyloidogenic Core Domains Systematic studies of Ab analogues by electron microscopy and X‐ray diffraction show that the core domain of amyloid fibrils consists of short peptides (Fraser et al., 1991, 1992; Inouye et al., 1993; Kirschner et al., 1987), and a recent study of PHF tau shows that only three residues is sufficient (Goux et al., 2004). Similarly short sequences for core domain of various amyloid fibrils have been identified (Tables IV and V), including VYK for tau PHF (Goux et al., 2004), LVFF for Ab (Inouye and Kirschner, 1996), NFGSVQ for medin (Reches and Gazit, 2004), DFNKF for calcitonin (Reches et al., 2002; Tsai et al., 2005), and GNNQQNY (Balbirnie et al., 2001; Diaz‐Avalos et al., 2003) for yeast prion Sup35 (Wickner et al., 2000). The homopolymeric domains of poly(l‐alanine) (Shinchuk et al., 2005) and poly(l‐glutamine) peptides (Sharma et al., 2005) that contain charged flanking residues form crystalline b‐sheet structures, indicating that the flanking residues may not impact the folding of the core. This suggests that hydrogen bonding between the side chains and the peptide backbone of the core may stabilize the b‐sheet assembly. Our current understanding, based on the aforementioned studies, is that amyloid formation is promoted via the interaction between short b‐strands. These core domains often contain aromatic residues which form p–p interaction


199

Table IV Diseases Involving Amyloida Disease

Amyloidogenic protein

Creutzfeldt‐Jakob disease (CJD) Alzheimer’s disease (AD) Hemodialysis‐related amyloidosis Primary systemic amyloidosis Secondary systemic amyloidosis Familial amyloid polyneuropathy I Familial amyloid polyneuropathy III Cerebral amyloid angiopathy Finnish hereditary systemic amyloidosis Type II diabetes Injection‐localized amyloidosis Medullary thyroid carcinoma Atrial amyloidosis Nonneuropathic systemic amylodosis Hereditary renal amyloidosis Parkinson’s disease Huntington’s disease

Prion (PrP) Ab and tau b‐2 microglobulin Immunoglobulin light chain Serum amyloid A (SAA) Transthyretin (TTR) Apolipoprotein A‐1 Cystatin A Gelsolin Islet amyloid polypeptide (amylin) Insulin Calcitonin Atrial natriuretic factor Lysozyme Fibrinogen a‐Synuclein Huntingtin (polyglutamine)

a

Modified from Kelly (1996) and Sipe and Cohen (2000).

(Reches et al., 2002), Gln and Asn which can form a hydrogen‐bonding network (polar zipper) via their side chains (Perutz et al., 1994), and polyalanine which provide strong van der Waals contact in quarter‐ staggered arrangement (Arnott et al., 1967; Inouye et al., 2000; Nguyen et al., 1995; Shinchuk et al., 2005). Therefore, the initial event in amyloid formation is likely to involve intramolecular turn formation (Bond et al., 2003; Lazo et al., 2005) and intermolecular interactions between neighboring, short b‐strands. In the following, we present the examples of polyalanine and polyglutamine.

VI. Polyalanine A.

Polyalanine Forms Cross ‐b Structure

X‐ray diffraction reveals that the polypeptide chains in the fibrillar assemblies formed by polyA11, polyA13, and polyA20 are in the cross‐b arrangement, that is, where the b‐strands run perpendicular to the fiber direction (Fig. 8) (Shinchuk et al., 2005). The X‐ray patterns show broad,

200

Table V Oligopeptides That Constitute Core Structures in Various Amyloids Sequence

Protein

Structure

References Sharma et al. (2005) Shinchuk et al. (2005) Makin et al. (2005) Papanikolopoulou et al. (2005) Goux et al. (2004); Inouye et al. (2006)

Huntingtin PABPN1 Model Adenovirus fiber Tau

Reverse‐turn b‐sheet Quarter‐staggered b‐sheet Antiparallel b‐sheet b‐Sheet Tubular assembly of b‐sheets

NFGSVQ NHVTLSQ LVVPDGLFV; LFVPDALFV; LVVPDAibLVV; MLFVPDALVVF Ab25–35 A‐Aib‐V; A‐Aib‐I; AGV IIGLM; FIGLM FF GNNQQNY

Medin b2‐Microglobulin Model

b‐Turn b‐Hairpin

Ab Model Ab31–35 Model Yeast prion Sup35

b‐Sheet and b‐turn b‐Turn Reverse turn Tube b‐Sheet

Octapeptide LANFLV; FLVHSS KFFE; KVVE DFNK; DFNKF STVIIE FF; VV FGAIL; NFGAIL AGAAAAGA Aminocyclohexane carboxylic acid

Lanreotide Islet amyloid (IAPP) Model Calcitonin Model Model IAPP Prion Model

Tube

b‐Sheet Tube Slab Cyclic b‐chain

Reches and Gazit (2004) Ivanova et al. (2004) Aravinda et al. (2004) Shanmugam and Polavarapu (2004) Maji et al. (2004) Bond et al. (2003) Reches and Gazit (2003) Balbirnie et al. (2001); Diaz‐Avalos et al. (2003) Valery et al. (2003) Scrocchi et al. (2003) Tjernberg et al. (2002) Reches et al. (2002) Lopez De La Paz et al. (2002) Go¨rbitz (2001) Tenidis et al. (2000) Nguyen et al. (1995) Amorin et al. (2003)


Polyglutamine Polyalanine KFFEAAAKKFFE GAITIG VYK; IVYK; VQIVYK

201


0.5 (201) (211) (200)

Intensity

0.8 0.6

(002)

(201) (211) 0.4

* (002)

0.3

* (001)

0.4 0.2

A13

A7

Intensity

1

0

0.4 0.2 0.3 0.1 Reciprocal coordinate (1/Å)

0

0.1 0.2 0.3 Reciprocal coordinate (1/Å)

0.4

Fig. 8. X‐ray diffraction data and analysis for polyalanine assemblies. (Top) Observed intensities (arbitrary scale) for equatorial scatter showing the (001) and (002) reflections, meridional scatter showing the (200) reflection, and off‐meridional scatter with the (201) and (211) reflections; the unit cell is orthogonal. Low‐angle equatorial reflection indicated by *. (Bottom) Calculated electron density map with projection along the chain direction (b‐axis)—hydrogen‐bonding direction (a‐axis) is horizontal and intersheet direction (c‐axis) is vertical—for solubilized/dried polyAla7 and polyAla13. Lattice constants: polyAla7, a ¼ 9.61 A˚, b ¼ 6.91 A˚, c ¼ 11.33 A˚; polyAla13, a ¼ 9.57 A˚, b ¼ 7.02 A˚, c ¼ 10.74 A˚. All samples show the quarter‐staggered arrangement of b‐sheets (i.e., the neighboring sheets are shifted by one quarter of the unit cell distance) a along hydrogen‐bonding direction. See Shinchuk et al. (2005) for further details.

strong equatorial reflection at 5.4 A˚ and a pair of strong, off‐meridional reflections at 4.44 and 3.74 A˚. These three reflections are indexed as (002), (201), and (211) of an orthogonal unit cell having dimensions a ¼ 9.6 A˚, b ¼ 6.9 A˚, and c ¼ 10.8 A˚, where the a, b, and c axes are in the hydrogen‐ bonding, b‐strand, and intersheet directions. The absence of the (200) reflection indicates that the b‐sheets are offset with respect to one another by one‐quarter of the unit cell dimension (Inouye and Kirschner, 1998). This arrangement, originally discovered for Tussah and Bombyx mori silk

202


(Marsh et al., 1955a,b), shows that the side chains pack closely against one other. Previous studies on homopolymers of poly(l‐alanine) were based on analyses of X‐ray patterns from fibers that had been steam‐stretched, hot‐rolled, or steam‐pressed (reviewed by Fraser and MacRae, 1973). Samples formed this way yield extended b‐pleated sheets (Arnott et al., 1967) in which the b‐strands are parallel to the direction of stroking or stretching. By contrast, the gels formed by water‐soluble short polyalanine peptides show cross‐b assemblies (Shinchuk et al., 2005). This organization results in a strong equatorial (002) reflection at 5.4 A˚, corresponding to the intersheet spacing for alanine side chains (Brown and Trotter, 1956), and in off‐meridional arcs at about 4.44 and 3.74 A˚, corresponding to the (201) and (211) reflections for a silk‐like lattice having quarter‐staggered b‐sheets (Marsh et al., 1955a,b).

B.

Polyalanine Extension in Oculopharyngeal Muscular Dystrophy

The pathological hallmark of oculopharyngeal muscular dystrophy (OPMD) is the appearance of tubulofilamentous inclusions in the nuclei of skeletal muscle fibers (Ku¨hn and Wahle, 2004). The toxicity may arise directly from the oligomeric or protofilament aggregates or from sequestration of RNA by the inclusion body (Kuhn and Wahle, 2004). OPMD fibrils may be brought about or precipitated by intermolecular hydrogen bonding between polyalanine domains, and this interaction may be stronger for the mutant PABPN1 which contains a larger number of alanine residues. This scenario appears to be consistent with the formation of 120‐ to 140‐A˚‐diameter fibrils from the N‐terminal domain of wild‐type PABPN1 (N‐WT), which contains 10 alanine residues, and of the mutant protein containing an extra 7 alanines [N‐(þ7)Ala]. Fibrils do not form when the N‐domain lacks alanine residues (PABPN1‐Ala) (Scheuermann et al., 2003). Based on its amino acid sequence/composition data, two distinct domains in the nascent PABPN1 protein are predicted: one is the structurally irregular N‐terminal domain (residues 1–125); and the other is the (globular) a þ b C‐terminal domain (residues 126–306). In the fibril, the N‐terminal domain may become folded into a b‐sheet, while the C‐terminal domain may remain globular. This type of structural transformation is similar to that modeled for the prion rod (see above), as the normal cellular form of prion contains a flexibly disordered N‐terminal (residues 90–145) and a globular C‐terminal domain. Moreover, transition to the pathogenic form (scrapie) is initiated by b‐sheet formation within the N‐terminal polyalanine‐rich domain of A8A (AGAAAAGA).


VII. A.

203

Polyglutamine

Polyglutamine Forms b‐Crystallites But Not a b‐Helical Nanotube

Analysis of polyglutamine assemblies by X‐ray diffraction has yielded two different types of proposed structures: one that is a water‐filled nanotube (Perutz et al., 2002a) and one that is composed of b‐crystallites (Sharma et al., 2005; Sikorski and Atkins, 2005) (Figs. 9 and 10). Among the different polyglutamine peptides studied, an oriented pattern was recorded only for Asp2Gln15Lys2 (Perutz et al., 1994, 2002b; Sharma et al., 2005), and the fiber pattern shows b‐crystallites with a cylindrical axis along the b‐strand direction (equatorial). The unit cell is monoclinic, and the cell constants are a ¼ 9.6 A˚, b ¼ 7.2, c ¼ 8.4, and g¼ 93 (Sharma et al., 2005). The meridional reflections (at 8.4, 4.8, 4.2, 2.8 A˚) can be indexed two dimensionally as (h0l), and the equatorial ones (at 3.6, 3.2 A˚) as (020) and (021). In addition, there is an off‐meridional reflection at 3.9 A˚. PolyGln15 shows a slab‐like morphology, with 30‐A˚‐period stacking in the b‐direction. To account for this dimension, the peptide must be folded in a reverse turn in the middle of the 15‐residue b‐strand.

Fig. 9. X‐ray intensity distributions (arbitrary scale) from aggregates formed by different polyglutamine peptides (Qn, for n ¼ 8, 15, 28, 45): polyGln45 (dried), polyGln28 (vapor hydrated), polyGln15 (vapor hydrated), and polyGln8 (lyophilized). The vertical bars indicate the positions of the Bragg reflections. The first interference peak for slab stacking of Q8 is indicated by *. See Sharma et al. (2005) for further details.

204


Fig. 10. Electron density projection along b‐strand direction—hydrogen‐bonding direction (a‐axis) horizontal, and intersheet direction (c‐axis) vertical—and skeletal models of polyGln8 (Q8) and polyGln45 (Q45) assemblies. The unit cell for both peptides was monoclinic, with lattice constants a ¼ 9.73 A˚, b ¼ 7.14 A˚, c ¼ 8.16 A˚, and g ¼ 95.7 for Q8, and a ¼ 9.66 A˚, b ¼ 7.10 A˚, c ¼ 8.33 A˚, and g ¼ 94.0 for Q45. The side chains are nearly overlapped in the hydrogen‐bonding direction. This difference in side chain conformation and disorder likely accounts for the differences in observed intensity between their diffraction patterns.

Comparable powder patterns were recorded for polyGln8 and polyGln45 (Fig. 9) indicating that polyglutamines having different numbers of residues fold similarly into a b‐crystallite type of assembly (Sharma et al., 2005). From the size of the stacking period, polyGln45 must form multiple reverse turns. Whether the turn creates intrachain H bonding between the antiparallel b‐chains and/or an intrachain intersheet interaction has been controversial (Sharma et al., 2005). The H‐bonded b‐chains are antiparallel in the former and parallel in the latter (Kajava et al., 2004). In fact, these two types of turn formation may coexist, as we showed earlier for the possible conformational polymorphism of the H1–H2 domain of PrP (see Fig. 5 in Inouye and Kirschner, 1998). While a polar‐zipper model was initially proposed for Asp2Gln15Lys2 (Perutz et al., 1994), more recently a water‐filled nanotube was proposed in which the homopolymer in the b‐conformation forms a helical array having 20 residues per turn (Perutz et al., 2002a,b). (In the earlier work, the diffraction pattern had been interpreted as a fiber pattern—that is, with the 4.8‐A˚ reflection in the fibril direction.) Rather than being attributed to intersheet stacking, the 8.4‐A˚ reflection was not accounted for. Further,


205

the absence of a 10‐A˚ reflection was ascribed to the presence of only a single b‐sheet comprising the thin wall of the tube. This model reproduces the observed 4.8‐A˚ reflection and the low‐angle equatorial maximum at 30 A˚, but not the other reflections (e.g., meridional 4.2 and 2.8 A˚, off‐meridional 3.9 A˚, and equatorial 3.6 and 3.2 A˚).

B. Reverse Turn May Be Crucial for Fibril Formation It has been argued that cell death such as occurs in CAG extension diseases including Huntington’s is caused by the accumulation of insoluble aggregates consisting of polyglutamine nanotubes in which the polyGln stretches have >40 residues (Perutz et al., 2002a,b). However, polyglutamine stretches with residue numbers from 15 to 45 all form slab‐like b‐structures (Sharma et al., 2005). The specific pathological effect of polyGlnn for n > 40 may be related to the rapid accumulation of insoluble fibrils due to reverse‐turn formation. A similar correlation between the formation of reverse turns and the initial event in amyloid fibril formation has been suggested from an analysis of powder diffraction from Ab31–35 (Bond et al., 2003) and by crystallography of synthetic peptides containing D‐Pro (Aravinda et al., 2004).

VIII. Conclusions X‐ray fiber and powder diffraction show b‐crystallite structure for alanine‐rich domains of prion peptides and of polyalanine. The quarter‐ staggered arrangement of alanine residues likely enhances the stability of the assembly, and may be involved in PrPC–PrPSc heterodimeric interactions. Not all of the recorded X‐ray spacings from polyglutamine are accounted for by the b‐helical nanotube model, but they are by the b‐crystallite model, which is stabilized by polar–zipper interactions between glutamine residues. Similar interaction between glutamine and asparagine residues in prion may also be involved in PrPSc formation. Short sequences likely initiate the formation of b‐assemblies for various amyloid fibrils—for example, formation of an intramolecular turn or intermolecular interaction may allow proximal, short core segments to be involved in the initial event of amyloidogenesis. Reagents with potential inhibitory effects on this interaction, for example, Congo red (Caspi et al., 1998; Caughey and Race, 1992), porphyrins and phthalocyanines (Caughey et al., 1998), tetracycline (Forloni et al., 2001, 2002; Tagliavini et al., 2000), and acridine and phenothiazine (Korth et al., 2001), may be useful for therapeutic purposes.

206


Acknowledgments We thank our colleagues for their valuable contribution to this research: fiber diffraction— Jeremy Bond, Sean Deverin, Dr. Leonid Shinchuk, and Dr. Deepak Sharma; prions—Drs. Jack Nguyen, Michael Baldwin, Haydn Ball, Robert Fletterick, Fred Cohen, Stanley Prusiner, and Mario Salmona; polyalanine—Drs. Sylvie Blondelle and Natalia Reixach; and polyglutamine— Dr. Ron Wetzel. We also thank Dr. Andrew Bohm (Department of Biochemistry, Tufts University) for graciously granting us access to their X‐ray diffraction facility.

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FROM THE POLYMORPHISM OF AMYLOID FIBRILS TO THEIR ASSEMBLY MECHANISM AND CYTOTOXICITY By LAURENT KREPLAK AND UELI AEBI M.E. Mu¨ller Institute for Structural Biology, Biozentrum, University of Basel, CH‐4056, Basel, Switzerland

I. II. III. IV. V. VI.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Polymorphism of Amyloid Fibrils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Soluble Forms of Amyloid Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Depicting Intermediate Stages of Amyloid Fibril Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . What Is the Mechanism of Small Oligomer‐Induced Cytotoxicity? . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Extracellular amyloid deposits are present in a variety of diseases. They contain amyloid fibrils that arise from the association of proteins or peptides. At the molecular level, all these fibrils share a common assembly principle based on a conformational change of the protein precursor leading to the formation of a cross‐b sheet structure. The smallest observed fibrils in vitro, often called protofibrils, are 4–5 nm in diameter. An amyloid fibril is generally composed of several of these protofibrils and may adopt different morphologies such as ribbons, sheets, or multistranded cables. This polymorphism was observed with many different amyloid‐forming peptides and proteins using electron microscopy. The need to understand the molecular origin of this effect as well as the desire to find inhibitors of fibril formation has driven researchers toward the dissection of amyloid fibril assembly pathways. We review the current knowledge on amyloid polymorphism and discuss recent findings in the field concerning amyloid fibril assembly pathways and cytotoxicity mechanisms.

I.

Introduction

The term amyloidosis is associated with a heterogeneous group of protein‐folding disorders. These diseases, like type 2 diabetes and Alzheimer’s disease, are characterized by the presence of macroscopic abnormalities, amyloid deposits, in one or more organs of the patients. The proteins or protein fragments accumulated in these deposits belong to three main ADVANCES IN PROTEIN CHEMISTRY, Vol. 73 DOI: 10.1016/S0065-3233(06)73007-8

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classes (Sipe and Cohen, 2000): (i) immune response proteins such as immunoglobulin chains, (ii) endocrine hormones like insulin or the islet amyloid polypeptide (also called amylin), and (iii) transport molecules such as transthyretin. Amyloid deposits were first identified by Virchow in 1851 using iodine and sulfuric acid (Virchow, 1851). Later on, Congo red was used as a histochemical stain (Bennhold, 1922) in conjunction with polarization microscopy (Divry and Florkin, 1927). With this technique, the deposits are characterized by an intense birefringence that is in agreement with their fibrillar architecture at the molecular scale. However, the binding mechanism of Congo red to amyloid fibrils is still ill understood and other dyes have been introduced like thioflavin T (Goldsbury et al., 2000b; LeVine, 1993) for which descriptions of the binding mechanism at the molecular level are becoming available (Khurana et al., 2005; Krebs et al., 2005). The first description of amyloid fibrils came from the electron microscopy (EM) studies of Cohen and Calkins in 1959 (Cohen and Calkins, 1959; Shirahama and Cohen, 1967). Since then, many fibrils of diverse origins in humans and animals have been analyzed. They generally appear as rigid rods with a diameter of 5–10 nm and a length in the range 100 nm to several micrometers (Fig. 1A and B). X‐ray fiber diffraction patterns of the fibrils reveal two main characteristic reflections at 0.47 and 1 nm (Eanes and Glenner, 1968). These are strong indications for a cross‐b structure in which the polypeptide chains form b‐sheets with their constituent b‐strands running roughly perpendicular to the fibril axis (Fig. 1C and D; Sunde et al., 1997). Other structural methods like solid‐state nuclear magnetic resonance (NMR) (Benzinger et al., 1998; Petkova et al., 2002)

Fig. 1. Structure of amyloid fibrils formed by the human amylin peptide. Negatively stained (A) and metal shadowed (B) fibrils formed by human amylin (adapted from Goldsbury et al., 2000a). (C) A human amylin fibril model formed by three protofibrils having a superpleated b‐structure (adapted from Kajava et al., 2005). Only Ca traces of the polypeptide chains are shown. (D) Atomic model of the cross‐b motif formed by the human amylin peptide (adapted from Kajava et al., 2005). Scale bar, 100 nm (A and B).

AMYLOID FIBRIL ASSEMBLY PATHWAYS AND CYTOTOXICITY MECHANISMS

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and Fourier transform infrared spectroscopy (FTIR) (Hiramatsu et al., 2004) have confirmed that a b‐sheet structure is the basic element of the amyloid fibril. However, these data can be interpreted in a variety of ways and several distinct models have been proposed for the fibril architecture (Fig. 1C; Flock et al., 2006; Jimenez et al., 1999; Kajava et al., 2005; Wetzel, 2002). Unfortunately, the description of amyloid fibrils given above is simplistic since in vitro self‐assembly of amyloid peptides and proteins yields polymorphic structures, as has been commonly observed in the past for other protein assemblies such as actin filaments (Millonig et al., 1988) and intermediate filaments (Herrmann and Aebi, 1999). On the one hand, assembly polymorphism complicates the characterization of fibril structure. On the other hand, it offers some insight into fibril formation. For this reason a more rational understanding of amyloid fibril formation at the molecular level is a key issue in the field of amyloidosis. It was long thought that amyloid fibrils were the toxic element responsible for cell death in infected tissues (Lorenzo and Yankner, 1994) and this idea drove the search for inhibitors of fibril formation (Tomiyama et al., 1994). However, several studies have shown the inherent toxicity of small soluble oligomers of amyloid peptides (Hartley et al., 1999; Lambert et al., 1998) even in the case of nonpathogenic amyloid‐forming proteins like HypF and SH3 (Bucciantini et al., 2002). These two opposite explanations for the cell toxicity of amyloid deposits have prompted recent systematic studies of amyloid fibril assembly pathways. To pursue such studies, several groups, including ours, have employed scanning force microscopy (SFM) as a tool to monitor the assembly of single amyloid fibrils on solid supports (Blackley et al., 2000; Goldsbury et al., 1999; Hoyer et al., 2002). Such a real‐time visualization of the dynamic process of amyloid fibril assembly has offered the opportunity to better understand the molecular origin of the observed polymorphism (Goldsbury et al., 2005; Green et al., 2004a), and the effects of chemical compounds (Goldsbury et al., 2005) or specific mutations (Green et al., 2003) on amyloid fibril assembly. In this chapter, we first describe amyloid fibril polymorphism as seen by EM. We then show how SFM was used to depict intermediate stages of amyloid fibril assembly. Finally, we discuss the putative mechanism that might explain the cytotoxicity induced by these assembly intermediates.

II.

Polymorphism of Amyloid Fibrils

The most detailed accounts of amyloid fibril polymorphism have come from EM studies of four different proteins or peptides, namely amyloid‐b (Ab1–40 or Ab1–42), transthyretin, calcitonin, and human amylin. The fibrils

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Fig. 2. Electron micrographs highlighting the polymorphism of amyloid fibrils. (A) A single human calcitonin protofibril with a diameter of 4 nm (adapted from Bauer et al., 1995). (B) Different morphologies present in a transthyretin fibril preparation. Black arrowheads show oligomers of different sizes, the black arrow points to a 9‐ to 10‐ nm‐wide fibril, and the white arrowhead marks an 4‐nm‐wide fibril (adapted from Cardoso et al., 2002). (C–F) Human amylin fibril ribbons (adapted from Goldsbury et al., 1997). (C) A single 5‐nm‐wide protofibril. (D–F) Ribbons containing two (D), three (E), or five (F) 5‐nm‐wide protofibrils. (G) A twisted ribbon made of four 5‐nm‐wide protofibril subunits of Ab1–40 (adapted from Goldsbury et al., 2000b). Scale bar, 50 nm (A–G).

were assembled in vitro from synthesized or recombinantly expressed proteins in various buffer conditions. In all cases, fibrils of different diameters and morphologies were observed as well as larger assemblies. For calcitonin, the thinnest single fibril, the ‘‘protofibril,’’ had a diameter of 4–5 nm and was observed at low (0.1–1 mM) calcitonin concentration (Fig. 2A; Bauer et al., 1995). For transthyretin, short and flexible protofibrils 4–5 nm in diameter were observed, but the prominent species was an 8‐nm diameter and up to 300‐nm‐long fibril (Fig. 2B; Cardoso et al., 2002). For human amylin, 5‐nm‐wide protofibrils were rarely depicted by themselves (Fig. 2C), but they could be readily identified as a distinct building block of wider fibrils (Fig. 2D–F; Goldsbury et al., 1997). The wider fibrils


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were multistranded cables (Fig. 2D), flat ribbons (Fig. 2E), or twisted ribbons (Fig. 2F), with both the multistranded cables and the twisted ribbons having a left‐handed twist (Goldsbury et al., 1997). Multistranded cables and ribbons were also observed for calcitonin (Bauer et al., 1995). These appeared to contain laterally associated 8‐nm‐ wide fibrils, whereas the diameter of the individual strands within the multistranded cables could not be measured directly from the images. To estimate this diameter, the authors plotted the helical crossover spacing of the cables as a function of their diameter. Using a linear regression fit, the data were extrapolated to zero, yielding a width of 4.1 nm, similar to the width of the single protofibrils (Fig. 2A; Bauer et al., 1995). For Ab it is worth noting that while single 5‐nm protofibrils were rarely imaged by EM or SFM, the thinnest single fibrils had a diameter around 8–9 nm and were termed ‘‘protofibrils’’ by many researchers in the field (Goldsbury et al., 2005; Harper et al., 1997, 1999; Lambert et al., 1998; Nielsen et al., 1999; Walsh et al., 1997, 1999). Flat and twisted ribbons formed from 5‐nm‐wide subunits, as seen with human amylin, were also depicted for Ab1–40 (Fig. 2G; Goldsbury et al., 2000b, 2005). From these data the notion emerges that in vitro assembled amyloid fibrils are multistranded cables or ribbons with a strand diameter of 4–5 nm. This unified interpretation of amyloid fibril morphologies has been further strengthened by mass‐per‐length (MPL) measurements of unstained fibrils using scanning transmission electron microscopy (STEM). For human amylin and Ab1–40, peaks at 20, 30, and 40 kDa/nm were measured. The 20 kDa/nm peak correlated in both cases with a fibril diameter of 8 nm made of two strands as seen by negative‐staining EM. This is consistent with a 4‐ to 5‐nm protofibril with an MPL of 9–10 kDa/nm (Goldsbury et al., 1997, 2000b, 2005). Similar MPL estimates were obtained with Ab1–42 (Antzutkin et al., 2002; Petkova et al., 2005). For the transthyretin fibrils, two main MPL peaks were observed at 9.5 and 14 kDa/nm and two minor ones at 19.9 and 25.8 kDa/nm. These are consistent with the existence of a 4‐ to 5‐nm protofibril (Fig. 2B) with an MPL of 4.8 kDa/nm (Cardoso et al., 2002). For calcitonin, MPL data are only available for the 8‐nm‐wide fibrils, with a rather broad peak at 27 kDa/nm (Bauer et al., 1995). With this single value it was not possible to extract a reliable value for the MPL of the calcitonin protofibril. Even though for each peptide or protein the MPL value for the protofibril may be different, all the available data can be summarized in a fairly simple scheme. The basic subunit of all amyloid fibrils is a 4‐ to 5‐nm‐wide protofibril whose detailed molecular architecture is species dependent. All the other structures observed can be either described as ribbons, sheets, or multistranded cables of protofibrils (Fig. 3).

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Fig. 3. A generalized model of amyloid fibril polymorphism based on the formation of straight or coiled fibrils composed of several 4‐ to 5‐nm‐wide protofibril subunits. Notice that the flat ribbons containing several protofibril strands may twist (Fig. 2F and G) and may ultimately form tubes (Bauer et al., 1995).

Interestingly, the morphology of the fibrils can be defined by just a few residues, as recently demonstrated by SFM for fibrils formed from prion proteins of different species (Jones and Surewicz, 2005). It is also possible that fibrils assembled from the same peptide exhibit a single morphology by EM, but can still be divided into subcategories according to their 3D structure and dynamic properties. In a direct attempt to prove this statement, Heise and coworkers studied the polymorphism of a‐synuclein fibrils by combining solid‐state NMR and EM. Using the same fibrillization conditions, they reproducibly obtained fibril preparations that were indistinguishable by EM (see Fig. 3 in Heise et al., 2005), but that could be separated into two categories according to solid‐state NMR (Heise et al., 2005). In an indirect way, we have also obtained a similar result for Ab1–42 fibrils when we recently analyzed the efficiency of an antibody raised against Ab1–16 to solubilize Ab1–42 amyloid fibrils. Within a population of fibrils with an apparently unique morphology as seen by EM, the majority (80–90%) of the fibrils were broken into small aggregates by this antibody, whereas a minority (10–20%) resisted solubilization and were decorated by antibodies (Greferath, Nicolau and Aebi, unpublished observations).


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These three examples emphasize the idea that a complete description of amyloid fibril polymorphisms will only be achieved when 3D structures at atomic detail become available (Luhrs et al., 2005). Last but not least, since the various morphologies observed for amyloid fibrils are defining the end point of the assembly process, an essential need is to properly define the early stages of fibril formation, namely the oligomeric states of the peptide or protein in solution prior to assembly into fibrils.

III.

Soluble Forms of Amyloid Peptides

In the various amyloidoses, the amyloid deposits are generally composed of a single protein. A minority of these proteins, like lysozyme, transthyretin, and some immunoglobulin light chains, misfold and associate into deposits as full‐length polypeptides (Kelly, 1996). The majority of amyloid‐forming proteins, however, are endoproteolysed to yield distinct peptide fragments. In both cases, the driving force of fibrillogenesis is a conformational change of the monomeric polypeptide chain that leads to enrichment of b‐sheet secondary structures. In this context, the structural state of the precursor and the way it is handled prior to fibril formation will have a strong influence on the assembly pathway. In the case of peptides, the conformation in solution has been shown to be sensitive to the solvent conditions. For Ab1–40 and various fragments, these effects have been reviewed extensively by Serpell who compiled NMR, FTIR, and circular dichroism (CD) data (Serpell, 2000). Serpell observed that organic solvents promoted an a‐helical conformation whereas, in aqueous buffer or in water, a b‐sheet conformation predominated (Goldsbury et al., 2000a; Serpell, 2000). In order to have a well‐defined starting point for the assembly process, the peptides are usually kept lyophilized until being dissolved in an organic solvent like 1,1,1,3,3,3‐ hexafluoro‐2‐isopropanol (HFIP) where the peptides assume at least partially an a‐helical conformation (Hirota‐Nakaoka et al., 2003; Maiti et al., 2004; Serpell, 2000). In the case of human amylin, this approach was particularly successful for the identification and characterization of early assembly intermediates (Green et al., 2004a; Higham et al., 2000).

IV.

Depicting Intermediate Stages of Amyloid Fibril Assembly

Now that we have defined the starting point and the final products of the assembly process, the next step is to investigate the origin of amyloid fibril polymorphism through a detailed study of the assembly pathways

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in vitro. This can be achieved by the combination of standard techniques like EM, CD, size‐exclusion chromatography (SEC), dynamic light scattering and chromophore‐binding assays. The two last techniques are generally used to follow the kinetics of amyloid fibril assembly, whereas SEC and EM are used to separate and visualize, respectively, metastable oligomers present in solution. Such a combined approach yielded the discovery of an early forming species during the assembly of Ab amyloid fibrils. These were called ‘‘protofibrils’’ (Walsh et al., 1997). Similar structures were isolated for transthyretin (Lashuel et al., 1998) and a‐synuclein (Conway et al., 1998). SFM was used as a tool to identify and characterize distinct morphologies and time courses of Ab fibril formation over several days. This highlighted ‘‘protofibrils’’ as possible intermediate states of assembly (Harper et al., 1997). However, ‘‘protofibrils’’ may not be on‐pathway intermediates for all of the polymorphic fibrillar forms of Ab(Goldsbury et al., 2005). Other prefibrillar species were observed by SFM and other methods after the onset of the aggregation process. These include the globular aggregates of HypF‐N (Relini et al., 2004) and Ab (Lambert et al., 1998). However, SFM is not only a tool to characterize morphologies, it has the ability to follow, in real time, dynamic processes occurring on a solid surface immersed in buffer (Stolz et al., 2000). For human amylin and Ab, conditions were found for which fibrils could assemble on a mica surface within a few hours (Blackley et al., 2000; Goldsbury et al., 1999, 2001, 2005). The surface‐confined assembly pathway of amyloid fibrils was then characterized in detail at the single filament level. Protofibrils were observed growing on the surface. These were 2.4 0.8 nm in height for human amylin (Goldsbury et al., 1999; Fig. 4A) and 6 0.5 nm for Ab (Goldsbury et al., 2005; Fig. 4B). In the case of Ab, variously sized globular aggregates appeared first (Fig. 4B) and, from initial studies, these were suggested to fuse over time to yield protofibrils (Blackley et al., 2000). However, in a more recent higher resolution study, Goldsbury and coworkers showed that while these fast‐forming aggregates possibly serve as stable nuclei for protofibril assembly, they do not generally fuse with each other (Goldsbury et al., 2005). For both human amylin and Ab, the protofibril growth rates were similar at both ends (Goldsbury et al., 1999, 2005). In the case of amylin, a single protofibril would occasionally grow from the tip of a thicker fibril (Goldsbury et al., 1999). Amylin fibrils growing on mica were rather straight and exhibited various heights. They were compatible with the protofibril hypothesis of amyloid fibril polymorphism (Fig. 3), but no multistranded cables were present (Goldsbury et al., 1999). In contrast, coiled fibrils were often observed by SFM for fibrils assembled in solution prior to being adsorbed to mica (Jansen et al., 2005; Kad et al., 2003; Relini et al., 2004). In the case of


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Fig. 4. Time‐lapse SFM experiments revealing the growth of single protofibrils and mature fibrils on mica. (A) Human amylin protofibrils (adapted from Goldsbury et al., 1999). (B) Bidirectional growth of a single Ab protofibril (adapted from Goldsbury et al., 2005). (C) Unidirectional growth of a mature Ab fibril (adapted from Goldsbury et al., 2005). Scale bar, 200 nm (A, B, and C).

Ab it was possible to follow the growth of twisted ribbons, with a periodic twist of 80–130 nm, by depositing seeds on mica prior to the injection of a fresh peptide solution (Fig. 4C; Goldsbury et al., 2005). In the case of human amylin, it was even possible to observe by time‐lapse SFM how fibrils are formed from an oligomeric nucleus by initial growth in height from

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2 nm up to 6 nm followed by extensive elongation (Green et al., 2004a). These results indicated that a small oligomer containing 10–50 human amylin monomers is the seed of amyloid fibrils and that the number of protofibrils within a mature fibril is specified by the dimensions of the seed at the onset of elongation (Green et al., 2004a). This is valid for constrained assembly onto a surface, but in solution multistranded cables and twisted ribbons are observed as described above. Emerging data are indicating that an initial heterogeneous mixture of variously sized oligomers may explain the final polymorphism of mature fibrils (Goldsbury et al., 2005; Green et al., 2004a; Petkova et al., 2005). Evidently, the origin of coiling has to be sought in the molecular architecture of the fibrils and, in particular, in the general tendency of b‐sheet structures to twist (Chothia, 1983). The predominance of one morphology over another may be controlled by external parameters, such as pH in the case of a‐synuclein (Hoyer et al., 2002). In the case of human amylin and Ab our understanding of the diversity in amyloid fibril architecture is the result of a recursive process, since the early morphological observations were followed by assessment of the assembly pathway which in turn yielded a better understanding of fibril polymorphism. However, this structural knowledge is secondary compared to the discovery of small oligomers, globular oligomers, and early ‘‘protofibrils’’ that appear to be extremely cytotoxic (Hartley et al., 1999; Lambert et al., 1998; Walsh et al., 1999).

V.

What Is the Mechanism of Small Oligomer‐Induced Cytotoxicity?

Slowly but definitely it is now becoming accepted that low molecular weight oligomers of amyloid peptides and proteins appear to be more toxic than mature amyloid fibrils at least to neuronal cultures and related cell lines (Huff et al., 2003). Some even argue that the fibrils formed at an early stage of amyloid diseases, while serving as transient protection against the toxic oligomers, may also represent a reservoir from which small oligomers may be generated (Hardy and Selkoe, 2002). To understand the molecular mechanism responsible for the cytotoxicity of distinct oligomers of amyloid‐forming peptides and proteins, a number of studies have focused on the interaction between amyloid‐forming peptides and lipid membranes, especially by patch clamping. This is how Arispe and coworkers observed for the first time an ion channel activity of Ab1–40 when inserted into a model lipid bilayer (Arispe et al., 1993). Similar behaviors were described for other amyloid‐forming peptides and proteins like human amylin (Mirzabekov et al., 1996) and b2‐microglobulin (Hirakura and Kagan, 2001).


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For Ab1–40 and a‐synuclein, physicochemical studies documented that the interaction with a membrane was dependent on the presence of acidic lipids and involved a conformational change of the peptide yielding an increase in its a‐helical content (Davidson et al., 1998; Hertel et al., 1997; Terzi et al., 1995, 1997). Concerning the nature and structure of such amyloid peptide or protein channels, oligomers with annular morphologies have in fact been observed by EM for a‐synuclein (Lashuel et al., 2002) and equine lysozyme (Malisauskas et al., 2003) even in the absence of any lipids or membranes. Channel‐like structures have also been reconstituted in liposomes and observed by SFM for Ab1–40, Ab1–42, human amylin, a‐synuclein, ABri, ADan, and serum amyloid A (Fig. 5A; Lin et al., 2001; Quist et al., 2005). Doughnut‐shaped structures with a diameter of 10–12 nm and a central hole size of 1–2 nm (Fig. 5B) were imaged on top of lipid membranes (Quist et al., 2005). However, the radius of curvature of the SFM tips meant that it is not possible to say whether the pores were really traversing the lipid bilayer. Another difficulty with these results is that during the reconstitution process, the peptide can access the lipid membrane on both sides which may not be the case in vivo (Quist et al., 2005). One way to solve this problem is to incubate a peptide solution on a lipid bilayer adsorbed to a solid support. This was performed for human amylin that was first dissolved in HFIP and injected into a drop of aqueous solution sitting above a mica‐ supported lipid bilayer (Green et al., 2004b). As mentioned above, HFIP was used to inhibit the growth of fibrils in the stock solution. By repeatedly imaging a given area of the bilayer, it was possible to observe the morphological changes over time induced by the peptide (Green et al., 2004b). Very small defects appeared on the surface, but no defined pores could be observed (Fig. 5C; Green et al., 2004b). Other groups have produced data consistent with the generalized increase in membrane permeability rather than the production of specific pores (Demuro et al., 2005). Interestingly, Quist and coworkers have obtained similar images of small defects for human amylin reconstituted into liposomes, except that the peptide was clearly visible lining the contour of the holes (Fig. 5A; Quist et al., 2005). Hence, in the case of human amylin, the peptide may be able to permeabilize lipid membranes without actually forming proper ion channels. A reassessment of previous patch‐clamping studies using monodispersed solutions of oligomers from various amyloid‐forming peptides including human amylin confirmed that no channel‐type conductivity jumps could be detected. Instead, a dramatic steplike increase of conductivity was observed on oligomer addition (Kayed et al., 2004) which, in fact, correlated rather well with the SFM results presented by Green and coworkers (2004a).

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Fig. 5. Interaction of amyloid‐forming peptides with lipid membranes. (A) SFM images of different amyloid peptides and proteins reconstituted into liposomes that were adsorbed to mica. Doughnut‐shaped aggregates are visible in each case. Scale bars 100 nm (adapted from Quist et al., 2005, National Academy of Sciences, USA). (B) High‐ magnification SFM images of the channel‐like aggregates (adapted from Quist et al., 2005, National Academy of Sciences, USA). Abstands for amyloid‐b and SAA stands for human aposerum amyloid A. (C) Time‐lapse SFM experiment displaying the effect of the human amylin peptide on a lipid bilayer that was adsorbed to mica (adapted from Green et al., 2004b). The arrows point to large bilayer defects that were present before peptide injection. The asterisks indicate defects that were created on peptide addition.

The controversy between the pore‐forming hypothesis and the permeabilization hypothesis is not resolved yet. One problem comes from the fact that current imaging and spectroscopic measurements cannot be performed on one and the same sample. This issue may soon be solved with the development of setups combining SFM directly with fluorescence


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spectroscopy (Owen et al., 2006; Reichlin et al., 2005). With such a setup it should be possible in the near future to investigate the mechanism of amyloid oligomer cytotoxicity both on model membranes and on single cells.

VI.

Conclusions

During the past 20 years, the study of amyloid fibrils has been very fruitful yielding detailed knowledge of their structure, assembly, and cytotoxicity mechanism. Interestingly, the main discoveries occurred in a fairly short period of time and are now giving us a clearer picture of amyloid fibril formation. Considering the complex self‐assembly mechanism that leads to fibril formation, our current understanding of this process is an impressive cross‐disciplinary achievement involving many different techniques and experimental approaches. However, since amyloid fibril polymorphism can be considered as only the ‘‘tip of the iceberg’’ in our understanding of the assembly process, its study is just a necessary prerequisite to eventually unravel the molecular sequelae yielding amyloid‐related pathologies. At this stage, the main unresolved issue concerns the molecular mechanisms leading to cytotoxicity. Describing the main players and the pathways involved is the major task ahead of us. Without any doubt, the search for therapeutic routes to cure amyloidoses will greatly profit from these studies.

Acknowledgments L. K. was supported by a grant from the Swiss Society for Research on Muscular Diseases awarded to U.A. and Sergei Strelkov. This work was also supported by grants from the NCCR ‘‘Nanoscale Science,’’ the Swiss National Science Foundation, and by the M. E. Mu¨ller Foundation.

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STRUCTURAL MODELS OF AMYLOID‐LIKE FIBRILS By REBECCA NELSON AND DAVID EISENBERG Howard Hughes Medical Institute, UCLA‐DOE Institute for Genomics and Proteomics, UCLA, Los Angeles, California 90095

I. II.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Refolding Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Insulin. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. SH3 Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Myoglobin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Prion Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Gain‐of‐Interaction Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Type I—Direct‐Stacking Models: Transthyretin and Superoxide Dismutase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Type II—Cross‐b Spine Models: GNNQQNY and b2‐Microglobulin . . . C. Type III—3D Domain‐Swapping Models: Cystatin C . . . . . . . . . . . . . . . . . . D. Type IV—3D Domain Swapping with a Cross‐b Spine: Ribonuclease A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Models of Natively Disordered Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Cylindrical b‐Helices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Other Models for Polyglutamine. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. HET‐s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Ure2p . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Amyloid b . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Fibril Properties and Their Relation to Structural Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Fibrillar Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Protofilament Substructure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Variable Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Cross‐b Diffraction Pattern . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Increase in b‐Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Congo Red Binding and Birefringence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . G. Self‐Association . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . H. Cooperative Kinetics of Formation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I. Extreme Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Amyloid fibrils are elongated, insoluble protein aggregates deposited in vivo in amyloid diseases, and amyloid‐like fibrils are formed in vitro from soluble proteins. Both of these groups of fibrils, despite differences in the sequence and native structure of their component proteins, share common ADVANCES IN PROTEIN CHEMISTRY, Vol. 73 DOI: 10.1016/S0065-3233(06)73008-X

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properties, including their core structure. Multiple models have been proposed for the common core structure, but in most cases, atomic‐level structural details have yet to be determined. Here we review several structural models proposed for amyloid and amyloid‐like fibrils and relate features of these models to the common fibril properties. We divide models into three classes: Refolding, Gain‐of‐Interaction, and Natively Disordered. The Refolding models propose structurally distinct native and fibrillar states and suggest that backbone interactions drive fibril formation. In contrast, the Gain‐of‐Interaction models propose a largely native‐like structure for the protein in the fibril and highlight the importance of specific sequences in fibril formation. The Natively Disordered models have aspects in common with both Refolding and Gain‐of‐Interaction models. While each class of model suggests explanations for some of the common fibril properties, and some models, such as Gain‐of‐Interaction models with a cross‐b spine, fit a wider range of properties than others, no one class provides a complete explanation for all amyloid fibril behavior.

I. Introduction Amyloid and amyloid‐like fibrils are elongated, insoluble protein aggregates deposited in vivo in amyloid diseases or formed in vitro from soluble proteins, respectively. Some 25 different proteins are known to be deposited as amyloid fibrils in humans (Westermark, 2005; Westermark et al., 2005). Despite a lack of sequence and structural similarity among fibril‐forming proteins, amyloid and amyloid‐like fibrils share common properties. These include an elongated, unbranched morphology; binding of the dye Congo red to produce a signature green birefringence under cross‐polarized light; and a cross‐b conformation defined by a characteristic X‐ray fiber diffraction pattern (Westermark et al., 2002). Although amyloid deposits began to be characterized in the 1850s (Cohen, 1986; Virchow, 1854), many of the morphological and structural similarities of amyloid fibrils from different sources became apparent only in the mid‐twentieth century. These included the birefringence of Congo red on binding amyloid deposits (Divry and Florkin, 1927; Missmahl and Hartwig, 1953), which has since been used as a diagnostic for amyloid. Electron microscopic (EM) examinations by Cohen and Calkins (1959) of amyloid deposits revealed long, unbranched, ‘‘delicate, wavy’’ fibrils with diameters ranging from 50 to 140 A˚. Fibrils also appeared to be composed of several, finer ‘‘protofilaments’’ (Gueft and Ghidoni, 1963; Serpell et al., 2000b; Shirahama and Cohen, 1967; Shirahama et al., 1973). The fibrillar morphology and green birefringence of Congo red suggested an ordered,

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parallel arrangement of linear substructure in the amyloid deposits (Puchtler et al., 1962; Wolman and Bubis, 1965). The ordered nature of amyloid fibrils was further demonstrated by Eanes and Glenner (1968) in their X‐ray fiber diffraction experiments. Amyloid fibrils give a cross‐b diffraction pattern, with perpendicular reflections at 4.7 A˚ (along the fiber axis or meridional direction) and 10 A˚ (equatorial direction) (Fig. 1A). This pattern was first reported by Astbury et al. (1935) for diffraction from ‘‘poached,’’ stretched egg albumin, which today would be termed an amyloid‐like substance. The pattern suggested that the albumin chains pack in an extended (i.e., b) conformation, with the chains perpendicular to the ‘‘axis of extension,’’ or fibril axis (Fig. 1B). The 4.7‐A˚ reflection corresponds to the 4.7‐A˚ stacking of b‐sheet strands, with hydrogen bonding parallel to the fibril axis. The 10‐A˚ reflection corresponds to the spacing of b‐sheets, with the sheet‐to‐sheet packing of side chains determining the exact packing distance (Fa¨ndrich and Dobson, 2002). Because this cross‐b pattern was found for all amyloid fibrils tested, Eanes and Glenner (1968) suggested that the fibrils have a common, cross‐b, core structure.

Fig. 1. Cross‐b structure of amyloid fibrils. (A) Cartoon representation of a cross‐b X‐ray diffraction pattern. The defining features are a meridional reflection at 4.7 A˚ and an equatorial reflection on the order of 10 A˚. The 4.7‐A˚ reflection is generally much brighter and sharper than the reflection at 10 A˚. (B) The cross‐b core structure of amyloid fibrils. Parallel b‐sheets are depicted, but the structure could equivalently be composed of antiparallel b‐sheets or a mix of parallel and antiparallel. The 4.7‐A˚ spacing of b‐strands within each b‐sheet is parallel to the long fibril axis. The depicted 10‐A˚ sheet‐to‐sheet spacing actually ranges from about 5 to 14 A˚ (Fa¨ndrich and Dobson, 2002), depending on the size and packing of amino acid side chains. Amyloid fibrils have diameters on the order of 100 A˚.

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The notion of a common core structure has been further supported by synchrotron X‐ray fiber diffraction patterns of several amyloid fibrils; the patterns show common reflections in addition to those at 4.7 and 10 A˚ (Sunde et al., 1997). Although these data give some insight into the arrangement of the amyloid fibril core, the exact molecular structure and organization of the proteins making up this common core have yet to be uniquely defined. The inherently noncrystalline, insoluble nature of the fibrils makes their structures difficult to study via traditional techniques of X‐ray crystallography and solution NMR. An impressive breadth of biochemical and biophysical techniques has therefore been employed to illuminate additional features of amyloid fibril structure. The resultant data have led to the proposal of numerous molecular models of amyloid fibril structure (Makin and Serpell, 2005). These models can be separated into three general classes (Fig. 2): (1) the ‘‘Refolding’’ models,

Model class

Native protein

Intermediate

Fibril

Refolding E.g., insulin, SH3, myoglobin, PrP

Gain-ofInteraction E.g., TTR, SOD, b2m, cystatin C, polyQ-RNase A

Natively Disordered E.g., polyQ, HET-s, Ure2p, Ab

?

Fig. 2. Classes of structural models of amyloid‐like fibrils. The Refolding models propose that a native protein (circle) partially or completely unfolds to attain a new fold (rectangle) in the fibril (stack of rectangles). In contrast, the Gain‐of‐Interaction models propose that only part of the native protein changes and takes on a new structure in the fibril. The remainder of the protein (partial circle) retains its native structure. The Natively Disordered models begin with disordered proteins or protein fragments, and these become ordered in the fibril. ‘‘PolyQ’’ refers to polyglutamine.


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in which a large fraction of the protein backbone differs in conformation between the native and fibrillar forms; (2) the ‘‘Gain‐of‐Interaction’’ models (Elam et al., 2003), in which only a small portion of the protein differs in conformation between the native and fibrillar states; and (3) the ‘‘Natively Disordered’’ models, composed of protein sequences or sequence fragments whose native structure seems disordered. In this chapter, we examine several examples of each class of model, and we consider the consistency of each model with experimental data.

II.

Refolding Models

Refolding models propose that the fibril‐forming protein exists in two distinct states: the native state and the fibrillar state (Fig. 2). In converting from one to the other, the protein must unfold, then refold. As the fibrillar state is common to proteins with dissimilar sequences, Fa¨ndrich et al. (2001) have suggested that refolding into fibrils is dominated by backbone interactions, which are available to all protein sequences. In this section, we discuss the Refolding models proposed for insulin, SH3 domain, myoglobin, and prion protein (PrP).

A. Insulin A prime example of a Refolding model is that of the insulin protofilament (Jimeńez et al., 2002). Insulin is a polypeptide hormone composed of two peptide chains of mainly a‐helical secondary structure (Fig. 3A; Adams et al., 1969). Its chains (21‐ and 30‐amino acids long) are held together by 3 disulfide bonds, 2 interchain and 1 intrachain (Sanger, 1959). These bonds remain intact in the insulin amyloid fibrils of patients with injection amyloidosis (Dische et al., 1988). Fourier transform infrared (FTIR) and circular dichroic (CD) spectroscopy indicate that a conversion to b‐structure accompanies insulin fibril formation (Bouchard et al., 2000). The fibrils also give a cross‐b diffraction pattern (Burke and Rougvie, 1972). Jimeńez et al. (2002) proposed a molecular model for the insulin protofilament based on these data and on electron cryomicroscopy (cryo‐EM) reconstructions of insulin fibrils. The fibrils show a number of twisted morphologies that seem to be alternative packings of similar protofilaments. The protofilaments have cross sections of 3040 A˚. The authors suggest a complete conversion to b‐structure and model the amyloid monomer as having four b‐strands (Fig. 3B). Each insulin chain contributes two of these b‐strands, and the chains align in a parallel stack, constrained by the interchain disulfide bonds. One pair of stacked b‐strands is curved

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Fig. 3. Refolding model of insulin protofilaments, from Jimeńez et al. (2002). (A) Ribbon diagram of the crystal structure of porcine insulin (PDB ID code 3INS), generated with Pymol (DeLano, 2002). The two chains are shown as dark and light gray with N‐ and C‐termini indicated. The dotted lines represent the three disulfide bonds: 1 is the intrachain and 2 and 3 are the interchain bonds. (B) Cartoon representation of the structure of monomeric insulin in the fibril, as proposed by Jimeńez et al. (2002). The thick, arrowed lines represent b‐strands, and thinner lines show the remaining sequence. The disulfide bonds are as represented in panel A, and N‐ and C‐termini are indicated. (Components of this panel are not to scale.) (C) Cartoon representation of an insulin protofilament, showing a monomer inside. The monomers are proposed to stack with a slight twist to form two continuous b‐sheets. (Components of this panel, including the protofilament twist, are not to scale.) In the fibril cross sections presented by Jimeńez et al. (2002), two, four, or six protofilaments are proposed to associate to form the amyloid‐like fibrils.

because of the steric constraints imposed by the intrachain disulfide bond. As the monomers stack to form extended sheets, a left‐handed twist of 1.5 –2.5 between strands recapitulates the twist of the fibril (Fig. 3C). This allows the protofilaments to interact via a single, common interface along the length of the fibril. Although this model fits the constraints imposed by the cryo‐EM data, it is unclear that the packed, curved, and flat sheets would yield a cross‐b diffraction pattern. The apparent shift from helical structure in the native state to b‐structure in the fibrillar state classifies this model as one of complete refolding.


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B. SH3 Domain A similar cryo‐EM study revealed structural features of fibrils of the SH3 domain of the p85a subunit of bovine phosphatidylinositol‐30 ‐kinase (Jimeńez et al., 1999). This 86‐amino acid SH3 domain natively folds into a b‐sandwich composed of 5 strands, 3 antiparallel strands in 1 sheet arranged perpendicular to 2 antiparallel strands in the facing sheet (Fig. 4A) (Booker et al., 1993). In vitro, the domain forms fibrils of multiple morphologies, including flat and twisted ribbons, and smooth and twisted tubular fibrils (Jimeńez et al., 1999). The fibrils give an X‐ray diffraction pattern with a strong 4.7‐A˚ and a weak 9.4‐A˚ reflection, indicating b‐structure (Guijarro et al., 1998). The cryo‐EM reconstructions of the twisted fibrils suggest a flattened cross section, composed of two pairs of protofilaments wrapped around a hollow center (Fig. 4B; Jimeńez et al., 1999). The protofilament cross sections measure 20 A˚ in the narrow dimension, which the authors suggest is room enough for two flat b‐sheets, but not for the native SH3 structure. The authors therefore suggest a model in which the five strands of the native SH3 domain rearrange to contribute to one or more of the fibril’s flat sheets (Fig. 4C). Each protofilament thus contains a pair of b‐sheets in the cross‐b conformation. This cryo‐EM‐based model for SH3 fibrils fits with the Refolding models, as it proposes a significant rearrangement of the native structure in forming amyloid‐like fibrils ( Jimeńez et al., 1999).

C.

Myoglobin

Although a detailed structural model has not been proposed for myoglobin fibrils, the protein was suggested to refold to form fibrils (Fa¨ndrich et al., 2001). Native myoglobin is largely a‐helical (Kendrew et al., 1960). In converting to fibril form, it seems to pass through an unfolded state, as monitored by CD spectroscopy and tryptophan fluorescence, forming fibrils suggested by FTIR spectroscopy to contain 35% b‐sheet and 11% a‐helix (Fa¨ndrich et al., 2003). The apparent transition from mainly a‐structure in the native state to largely b‐structure in the fibril via a possibly unfolded intermediate fits with the Refolding models.

D. Prion Protein The prion protein, implicated in diseases such as mad cow and Creutzfeldt‐ Jakob, is another that has been proposed to undergo extensive refolding to form fibrils. In its native, cellular conformation (PrPC), residues 23–124 are

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Fig. 4. Refolding model of SH3 domain fibrils, from Jimeńez et al. (1999). (A) Ribbon diagram of the solution structure of the SH3 domain of bovine phosphatidylinositol‐30 ‐kinase (PDB ID code 1PNJ), generated with Pymol (DeLano, 2002). The two sheets are shown as light and dark gray, and N‐ and C‐termini are indicated. (B) Cartoon representation of the cross section of an SH3 fibril, interpreted from Figs. 3C, D, and 5C of Jimeńez et al. (1999). The gray region corresponds to the density of the fibril as observed by cryo‐EM; the center appears hollow. The dotted circles outline the higher‐ density regions, corresponding to four suggested protofilaments. Each protofilament is proposed to be composed of two flat b‐sheets, represented by white rectangles. The authors note that extra loop regions could pack into the surrounding, lower‐density regions of the fibril. (C) Side view of a section of the fibril, interpreted from Fig. 5D of Jimeńez et al. (1999), showing the suggested protofilaments as gray cylinders and b‐strands as white arrows. The direction of the arrows does not correspond to chain direction. The figure shows how the b‐strands of SH3 might stack into flat b‐sheets and pack into protofilaments. This representation does not show the observed twist of the fibrils.


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flexible and disordered, and residues 125–230 form a structured, largely a‐helical domain, in which a‐helices 2 and 3 are linked by a disulfide bond (Fig. 5A; Zahn et al., 2000). Proteinase K treatment of the infectious PrPSc conformer followed by purification produces infectious prion rods composed of a fragment lacking the N‐terminal 89 residues (PrP 27–30) (McKinley et al., 1991; Prusiner et al., 1983). Prion rods in sucrose solution give an X‐ray diffraction pattern showing a weak 4.7‐A˚ and a diffuse 8.8‐A˚ reflection, suggested to indicate the presence of b‐structure (Nguyen et al., 1995). Optical spectroscopic measurements also suggest that PrPSc and PrP 27–30 have significant b‐sheet content, much higher than that of PrPC (Caughey et al., 1991; Gasset et al., 1993; Pan et al., 1993; Safar et al., 1993). Govaerts et al. (2004) proposed a parallel b‐helix model for prion rods that is consistent in overall dimensions with their low‐resolution EM studies of two‐dimensional PrP 27–30 crystals (Wille et al., 2002). In this model, residues 89–174 form 4 ‘‘coils,’’ or complete helical turns ( Jenkins and Pickersgill, 2001), of a left‐handed, parallel b‐helix (Fig. 5B). The coils of one monomer are proposed to stack on the coils of another to form an extended triangular b‐structure. Three of these triangular units pack together to form the fibril (Fig. 5C and D). The C‐terminal a‐helices (a2 and a3) of monomeric PrP are proposed to retain their native structure in the fibril and pack around the outside of the trimer (Fig. 5C and D). The presence of these helices in the prion rods is consistent with antibody binding studies (Peretz et al., 1997), the presence of a disulfide bond (Turk et al., 1988), and FTIR measurements (Wille et al., 1996). The parallel b‐helix model does not provide an 8.8‐A˚ sheet‐to‐sheet spacing, suggested by the 8.8‐A˚ reflection of the prion rod diffraction pattern (Nguyen et al., 1995). However, the authors (Govaerts et al., 2004) note that a number of b‐sandwich folds are consistent with the general structural requirements for a model of fibrillar PrP, although they do not attempt to model it as such. The proposed changes in conformation of residues 89–174 have led us to place this model (Govaerts et al., 2004) in the Refolding class, although the native disorder of residues 89–124 could equally place this model into the Natively Disordered class.

III.

Gain‐of‐Interaction Models

The ‘‘Gain‐of‐Interaction’’ model of fibril formation (Elam et al., 2003) proposes that a conformational change in a limited region of the native protein exposes a normally inaccessible interaction surface that drives fibril formation. In these models, the bulk of the protein retains its native

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Fig. 5. Refolding model of prion rods, from Govaerts et al. (2004). (A) Ribbon diagram of the solution structure of human PrP (PDB ID code 1QM0), generated with Pymol (DeLano, 2002). Residues 125–228 are shown. The structure is colored to show the boundary of the proposed refolding: the dark gray N‐terminal region changes structure, while the light C‐terminal region retains its native structure. The dotted line represents the disulfide bond, and N‐ and C‐termini are indicated. (B) Ribbon diagram of four coils of the left‐handed parallel b‐helix of UDP N‐acetylglucosamine O ‐acyltransferase from Escherichia coli (PDB ID code 1LXA), generated with Pymol (DeLano, 2002). This structure is similar to that proposed by Govaerts et al. (2004) for the refolded portion of the PrP prion rod. Note the loops emanating from the core b‐helix; these are common in parallel b‐helices. The PrP prion rod model also proposes two loops extend from the core helix. N‐ and C‐termini are indicated. (C) Cartoon representation of the PrP prion rod model (Govaerts et al., 2004), viewed down the fiber axis. The gray triangles represent the left‐handed parallel b‐helices, and the white rods represent the native helices a2 and a3. (D) Cartoon representation of the PrP prion rod model viewed approximately perpendicular to the fibril axis (based on Fig. 4B of Govaerts et al., 2004). Representations are the same as in panel C.


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Fig. 6. Gain‐of‐Interaction models. Type I, or the direct stacking models, propose that the native protein undergoes limited structural changes, forming two interaction surfaces that cause the protein to stack and form the fibril core. Type II, or the cross‐b spine models, propose that a small segment separates from the native protein and interacts with similar segments from other proteins to form the cross‐b‐structured core of the fibril. In the cartoon, the zigzagging lines represent the b‐strands, and the dotted lines represent the interstrand hydrogen bonds. Type III, or the 3D domain‐swapping models, propose that 3D domain swapping of monomers forms the fibril. Type IV, or the 3D domain swapping with cross‐b spine models, propose that monomers domain swap and the hinge loops stack into the cross‐b core of the fibril.

structure (Fig. 2). Gain‐of‐Interaction models are therefore in keeping with Anfinsen’s ‘‘thermodynamic hypothesis’’ (Anfinsen, 1973) that a protein’s sequence determines its native, lowest energy structure. Refolding models, which postulate two distinctly different structures—native and fibrillar— are not so easily reconciled with this long‐standing pillar of protein science. We divide the Gain‐of‐Interaction models into four types (Fig. 6). Type I are the direct‐stacking models, in which a small conformational change allows fibril formation via stacking of the subunits. Examples include fibril models of transthyretin (TTR) and superoxide dismutase (SOD). Type II models contain a cross‐b spine, with remaining portions of the protein packed around the spine. Examples include fibril models of the GNNQQNY peptide of Sup35p and b2‐microglobulin. Type III models form via three‐ dimensional (3D) domain swapping and include models of cystatin C fibrils. Type IV models contain 3D domain swapping and a cross‐b spine, as proposed for fibrils of a polyglutamine insertion mutant of ribonuclease A.

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A.

Type I—Direct‐Stacking Models: Transthyretin and Superoxide Dismutase

Transthyretin is a 147‐residue transporter protein that is deposited as amyloid in senile systemic amyloidosis and familial amyloid polyneuropathy (Benson and Uemichi, 1996; Saraiva, 1995; Westermark et al., 1990). The protein’s native structure is a 55‐kDa dimer of dimers, or homotetramer, composed mainly of b‐sheets (Blake et al., 1978). The native monomer‐ to‐monomer interface is formed by mutual, antiparallel extension of each monomer’s two sheets (Fig. 7A); both four‐stranded sheets in the monomers (DAGH and CBEF) become eight‐stranded sheets in the dimer (DAGHH0 G0 A0 D0 and CBEFF0 E0 B0 C0 ) (Blake et al., 1978). In vitro at low A

B C⬘ B⬘ D⬘

F⬘ H⬘ F

H G

C

E

A C

N

D

B

F E B

N A⬘

E⬘

B⬘ E⬘ F⬘

G⬘

C

B⬘ E⬘ F⬘ F E B

Fibril axis

B⬘ E⬘ F⬘ F E B Native interface Gain-of-Interaction interface

Fig. 7. Direct‐stacking Gain‐of‐Interaction model of TTR fibrils, from Serag et al. (2002) and Olofsson et al. (2004). (A) Ribbon diagram of a dimer of human TTR (PDB ID code 2PAB), generated with Pymol (DeLano, 2002). One monomer is colored yellow and the other green, and N‐ and C‐termini are indicated. The red strands and loops are those proposed to change conformation in the direct‐stacking model of the TTR fibril (Olofsson et al., 2004; Serag et al., 2002). (B) Cartoon representation of the direct‐stacking model of the TTR fibril (based on Fig. 4B of Serag et al., 2002). Colored rectangles represent strands and/or protein segments of the same color and name as shown in panel A. Open circles represent the native monomer‐to‐monomer interface, thought to be maintained in the fibril. Closed circles represent the new interface in the fibril, which creates a continuous b‐sheet.


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pH, TTR dissociates to form monomeric, dimeric, and higher‐oligomeric intermediates that can aggregate into fibrils (Lai et al., 1996; Lashuel et al., 1998; Olofsson et al., 2001). TTR fibrils give a cross‐b diffraction pattern (Blake and Serpell, 1996), and are suggested by CD spectroscopy to contain a lower b‐sheet content than the native tetramer (Olofsson et al., 2001). Based on site‐directed spin labeling and cross‐linking studies, Serag et al. (2001, 2002) proposed a ‘‘head‐to‐head, tail‐to‐tail’’ stack of TTR monomers in the fibril (Fig. 7B). The authors suggest that the native monomer‐ to‐monomer interface is retained in the fibril (Serag et al., 2001), providing a head‐to‐head interaction. For the tail‐to‐tail interaction, they suggest that the C strand, colored red in Fig. 7, moves out of the way, allowing B strands from separate dimers to interact in an antiparallel arrangement (Serag et al., 2002). A similar model was proposed by Olofsson et al. (2004) based on hydrogen/deuterium exchange experiments. The Olofsson model adds that the whole loop containing the C and D strands moves away to additionally expose the A strand in the gained interface (Olofsson et al., 2004). It is noteworthy that the core repeating unit of this model, that is, the two sheets AGHH0 G0 A0 and BEFF0 E0 B0 , fits the dimensions and twist proposed by Blake and Serpell (1996) for half of the protofilament core structure of TTR. Also, this model fits with the proposal that proteins have evolved mechanisms to prevent b‐strand aggregation (Richardson and Richardson, 2002), as the C and D strands seem to protect native TTR from aggregation. This direct‐stacking model (Olofsson et al., 2004; Serag et al., 2002) therefore proposes that TTR maintains much of its native structure, including the native dimer interface, in the fibrillar state. A new interaction interface is gained with the shifting of b‐strands at the ends of two sheets, driving fibril formation. A second direct‐stacking model of fibril structure was proposed by Elam et al. (2003) for human copper‐zinc SOD. SOD is a 32‐kDa, homodimeric, largely b‐sheet protein with antioxidant activity (Fig. 8A; Fridovich, 1989; Parge et al., 1992). Mutations in SOD are associated with familial cases of amyotrophic lateral sclerosis, also known as Lou Gehrig’s disease (Deng et al., 1993; Rosen et al., 1993). Elam et al. (2003) crystallized several SOD mutants and found filamentous arrangements of the dimers in the solved structures (Fig. 8B and C). In the structure of the S134N mutant, an altered loop conformation exposes the edges of two b‐strands, and these exposed edges serve as one interface for the gained interaction. The arrangement of dimers in the S134N mutant filament results in four of eight b‐strands per subunit sitting perpendicular to the long axis of the filament, with the sheet‐to‐sheet spacing approximately perpendicular to this axis (Fig. 8C). This arrangement is roughly consistent with a cross‐b structure, although a cross‐b diffraction pattern has not been reported. As SOD aggregates may

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Fig. 8. Direct‐stacking Gain‐of‐Interaction model of SOD filaments, from Elam et al. (2003). (A) Ribbon diagram of the crystal structure of human SOD (PDB ID code 1HL5). The two monomers of the dimer are colored in light and dark gray, their copper and zinc ions are shown as spheres, and disulfide bonds are represented by dotted lines. N‐ and C‐termini are indicated for one monomer. (B) Surface representation of three dimers of a crystalline filament of human SOD mutant S134N (PDB ID code 1OZU). (C) Ribbon diagram of the same structure and view as shown in panel B. The b‐strands arranged roughly perpendicular to the fibril axis are highlighted in black. All panels were generated with Pymol (DeLano, 2002).

belong to a class of aggregates distinct from amyloid (Westermark et al., 2002), it is difficult to interpret these results. Whether amyloid‐like or not, these SOD crystalline filaments are formed from a stack of native‐ like SOD subunits, in which a limited conformational change results in the gained interface.

B. Type II—Cross‐b Spine Models: GNNQQNY and b2‐Microglobulin A cross‐b spine structure consists of two or more flat or twisted b‐sheets, composed of parallel (Nelson et al., 2005) or antiparallel (Makin et al., 2005) b‐strands, in a cross‐b arrangement. The cross‐b spine model of fibril formation proposes that a short segment of the native protein changes conformation to form one or more b‐strands of a cross‐b spine. The seven‐ residue peptide GNNQQNY, derived from the prion‐determining domain


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Fig. 9. Cross‐b spine of GNNQQNY from the crystal structure of Nelson et al. (2005). (A) Pair of sheets of the GNNQQNY cross‐b spine, with backbones represented by arrows and side chains by ball‐and‐stick (PDB ID code 1YJP). The vertical half‐arrow shows the proposed fibril axis and indicates the twofold screw symmetry of the two sheets; a 180 rotation of one sheet about the axis, followed by a vertical translation of half the 4.9‐A˚ strand–strand spacing produces the exact positioning of the second sheet. The asparagine and glutamine side chains facing into the space between the two sheets pack to form a steric zipper. (B) Stick representation of the top view of the spine, looking down the fibril axis. The dark and light molecules correspond to the dark and light sheets of panel A. This view shows the interdigitation of the asparagine and glutamine side chains in the steric zipper. The position of the twofold screw axis is indicated by . (C) Cut‐away stick representation of a side view of the spine, looking along the strands. This view is from the left edge of panel A, with corresponding coloring. The view shows the stacking of glutamine (Q4 and 5) and asparagine (N3) side chains in the direction of the proposed fibril axis; hydrogen bonds between stacked side chains are shown as dotted lines. All panels were generated with Pymol (DeLano, 2002).

of the yeast prion Sup35p, forms amyloid‐like fibrils and crystallizes into a cross‐b spine structure, which has been determined to atomic resolution by X‐ray diffraction (Fig. 9; Balbirnie et al., 2001; Nelson et al., 2005).

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Fig. 10. Gain‐of‐Interaction models of b2m fibrils, from Ivanova et al. (2004) and Benyamini et al. (2003). (A) Ribbon diagram of the crystal structure of the human b2m monomer (PDB ID code 1LDS), generated with Pymol (DeLano, 2002). The dotted circle shows the location of the seven‐residue sequence proposed to be important in fibril formation (Ivanova et al., 2004), and N‐ and C‐termini are indicated. (B, C) Ribbon diagrams of the cross‐b spine model of hb2m fibrils of Ivanova et al. (2004); the two sheets are distinguished by light and dark gray coloring. (B) The cross‐b spine, made up of stacked b‐hairpins. The left and right views are related by a 90 rotation about the vertical fibril axis. The left view highlights the antiparallel stacking of hairpins, and the right view highlights the sheet‐to‐sheet spacing and 7 twist from strand to strand. (C) View of the cross section of the hb2m fibril, with the fibril axis pointing out of the page and slightly to the left. Two hairpins stack in each sheet, and the remainder of each native‐like hb2m monomer packs around the spine. N‐ and C‐termini are indicated.


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For this reason, the cross‐b spine model is on a more certain footing than most others discussed in this article. In this cross‐b spine, each GNNQQNY peptide assumes an extended b‐strand conformation, and these peptide strands stack parallel and in‐register to form flat b‐sheets (Fig. 9A). Two sheets pack together to form a steric zipper; the peptide side chains interdigitate to form a dry, tightly packed, highly self‐complementary interface (Fig. 9B). Additionally, the amide side chains in the interface hydrogen bond with their neighbors in the same sheet to form an extensive network of hydrogen bonds (Fig. 9C). The sum of these interactions provides stability and demonstrates that short protein segments may form a cross‐b core while the remainder of the protein retains its native structure. A cross‐b spine model was proposed for the fibril structure of human b2‐microglobulin (hb2m) (Ivanova et al., 2004). hb2m is a 99‐amino acid serum protein with a 7‐stranded b‐sandwich fold (Fig. 10A; Saper et al., 1991). In patients on long‐term kidney dialysis, the protein is deposited as amyloid fibrils in the joints (Floege and Ehlerding, 1996; Koch, 1992). In vitro ‐formed fibrils of hb2m give a cross‐b X‐ray diffraction pattern (Ivanova et al., 2004; Smith et al., 2003). Several studies have shown that segments of hb2m form amyloid‐like fibrils on their own (Ivanova et al., 2003; Jones et al., 2003; Kozhukh et al., 2002). Limited differences in the human (amyloid‐forming) and mouse (non‐ amyloid‐forming) b2m sequences led Ivanova et al. (2003, 2004) to examine the amyloidogenic properties of the C‐terminal region of b2m. A peptide spanning loop residues 83–89, of hb2m forms amyloid‐like fibrils on its own, whereas a peptide spanning the same region of the mouse b2m (mb2m) sequence does not (Ivanova et al., 2004). Substitution of the human peptide (83–89) sequence into the mouse protein confers fibril‐forming capability on mb2m, while substitution of the mouse peptide sequence into hb2m inhibits, but does not prevent, fibril formation. The authors (Ivanova et al., 2004) therefore propose a fibril model where residues 83–97, which make up the C‐terminal loop and strand of the native structure, form a b‐hairpin in the fibril. The b‐hairpins stack into antiparallel b‐sheets, and two sheets pack together at 11‐A˚ spacing to form the cross‐b spine (Fig. 10B). In this model, the rest of the protein retains its native structure, packing around and protecting the two b‐sheets of the spine (Fig. 10C). Thus, this model proposes the fibril forms when a new interaction is gained (D) Cartoon representation of the direct‐stacking model of hb2m fibrils, from Benyamini et al. (2003). Rectangles represent the intact strands, where b‐strands B, E, and D mostly obscure the view of strands F and C. b‐Strands A and G of the monomer have separated from the core (dotted lines), allowing stacking of the remaining strands to form a continuous b‐sheet. The gained interactions are indicated by the closed circles.

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through the folding and stacking of a C‐terminal b‐hairpin into a cross‐b spine. A different Gain‐of‐Interaction model, a direct‐stacking model, has also been suggested for the structure of the hb2m fibril (Benyamini et al., 2003). Benyamini et al. (2003) used computational protein docking methods to search for cross‐b fibrillar arrangements of native‐like hb2m. In the resulting model, the subunits of the fibril core are composed of b‐strands B through F, and they interact through parallel, backbone hydrogen bonding of strand B from one subunit to strand D of the next (Fig. 10D). The stacking of subunits into the fibril thus creates a continuous b‐sheet (strands BED‐BED‐BED. . .). The absence of strands A and G from the fibril core is supported by in vitro studies, including equilibrium denaturation monitored by NMR (McParland et al., 2002), H‐D exchange monitored by NMR (Hoshino et al., 2002), and limited proteolysis monitored by mass spectrometry (Monti et al., 2002). In summary, two different Gain‐of‐Interaction models have been proposed for the fibrillar structure of b2m. The cross‐b spine model (Ivanova et al., 2004) proposes a core composed of C‐terminal b‐hairpins, and the direct‐stacking model (Benyamini et al., 2003) proposes a core of native‐like b2m molecules with their N‐ and C‐terminal strands displaced.

C.

Type III—3D Domain‐Swapping Models: Cystatin C

3D domain swapping is a mechanism for protein homo‐oligomerization that has been proposed to be involved in amyloid fibril formation (Bennett et al., 1995; Schlunegger et al., 1997). Figure 11 illustrates 3D domain swapping. In the example of a 3D domain‐swapped dimer, two identical proteins exchange equivalent structural components, or domains, thereby dimerizing via a monomeric interface. This closed interface is the surface of interaction between the ‘‘swapped’’ domain and the rest of the protein. The closed interface is identical in the monomer and dimer (or higher oligomer), with the exception that in the oligomer, two separate polypeptide chains contribute to the interaction. Only one segment of the protein changes in conformation between the monomer and 3D domain‐swapped oligomer: the hinge loop. The hinge loop connects the swapped domain to the rest of the protein. The new, or gained, interface in the oligomer is termed the open interface, and if it provides a sufficiently favorable interaction, it can drive the equilibrium between monomer and oligomer toward the oligomeric state. 3D domain swapping shares several features with amyloid fibril formation. It is specific in that only one type of protein is contained in any given


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Fig. 11. 3D domain swapping. (A) A monomer that can undergo 3D domain swapping has two domains that are connected in sequence by the hinge loop. The domains interact through the closed interface. (B) In a 3D domain‐swapped dimer, the hinge loop changes conformation, the closed interfaces are re‐formed by domains from two separate molecules, and a new open interface is formed. The open interface may involve an interaction between the two hinge loops and may also incorporate surface regions of the native domains. (C) A run‐away 3D domain‐swapped oligomer is formed when monomers swap successively, leaving unpartnered domains at the ends of the linear structure.

oligomer. It is known to occur in a number of proteins that are unrelated in sequence and structure. And it requires partial unfolding of the subunit protein. An example of an open‐ended, linear, 3D domain‐swapped oligomer is shown in Fig. 11C. Also called a ‘‘run‐away’’ domain swap, it is one model for how 3D domain swapping could lead to amyloid formation (Bennett et al., 1995; Janowski et al., 2001; Klafki et al., 1993; Knaus et al., 2001; Liu et al., 2001; Sambashivan et al., 2005). 3D domain swapping may be involved in the deposition of fibrils in hereditary cystatin C amyloid angiopathy, a disease of young adults leading to fatal cerebral hemorrhage (Abrahamson, 1996; Olafsson and Grubb, 2000). Cystatins are cysteine protease inhibitors whose native fold consists of an a‐helix running across a five‐stranded, antiparallel b‐sheet (Fig. 12A; Bode et al., 1988). Several cystatins form inactive, 3D domain‐swapped dimers (Ekiel and Abrahamson, 1996; Jerala and Zerovnik, 1999; Zerovnik et al., 1997). In the dimer (Fig. 12B), the N‐terminal structural elements b1‐a1‐b2 of one subunit exchange positions with the same elements of the other subunit (Janowski et al., 2001). The loop (i.e., the hinge loop)

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Fig. 12. 3D domain swapping of cystatin. (A) Ribbon diagram of the crystal structure of chicken cystatin (PDB ID code 1CEW). The 108‐residue protein has two disulfide bonds, represented by dotted lines. The position of the hinge loop is indicated, as are the N‐ and C‐termini. (B) Ribbon diagrams of the crystal structure of a 3D domain‐ swapped dimer of human cystatin C (PDB ID code 1G96, Janowski et al., 2001), related by a 90 rotation about the horizontal. The 120‐amino acid monomers are colored as black and light gray to highlight the swapped domains. Human cystatin C also contains two disulfide bonds, which are not shown. The top panel shows the hinge loop, which has become a b‐strand connector between b2 and b3. N‐ and C‐termini are indicated. All panels were generated with Pymol (DeLano, 2002).

that had connected b2 and b3 in the monomer takes on an extended b‐conformation in the dimer, resulting in a continuation of b2 in one subunit into b3 of the other (Fig. 12B). Two models have been proposed for how this dimeric structure may relate to the structure of cystatin C in the fibril. The first ( Janowski et al., 2001) proposes that run‐away domain swapping (like that shown in Fig. 11C) can account for the assembly and stability of the fibril. In this model, one monomer would swap b1‐a1‐b2 into a second monomer, the second would swap its b1‐a1‐b2 into a third, and so on. The second model (Staniforth et al., 2001) proposes a direct stacking of domain‐swapped dimers, where b5 of each subunit of the dimer would interact with b1 of a subunit of the adjacent dimer. In this way, the dimers would stack to form continuous b‐sheets. Both models arrange the b‐sheets parallel to the fibril axis with component b‐strands perpendicular to the axis, as in a cross‐b structure, although no diffraction pattern has been reported for cystatin fibrils.


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A later study by Nilsson et al. (2004) examined the role of 3D domain swapping in cystatin C fibril formation. By engineering in a disulfide bond to prevent formation of the domain‐swapped dimer, the authors were able to reduce fibril formation by 80%. These results suggest that the 3D domain swapping seen in the dimer cannot uniquely account for cystatin C fibril formation; either fibril formation is achieved via a different mechanism or multiple pathways are possible, of which 3D domain swapping is one (Nilsson et al., 2004).

D. Type IV—3D Domain Swapping with a Cross‐b Spine: Ribonuclease A A Gain‐of‐Interaction model with swapped domains and a cross‐b spine was proposed for the polyglutamine insertion mutant of ribonuclease A (RNase A) (Sambashivan et al., 2005). RNase A is a 124‐residue enzyme of a‐ and b‐structure and contains four disulfide bonds (Fig. 13A) (Kartha et al., 1967). The protein forms 3D domain‐swapped dimers and higher‐ order oligomers on freeze‐drying from 40% acetic acid (Crestfield et al., 1963; Gotte et al., 1999; Liu et al., 2001, 2002). Insertion of a polyglutamine sequence (GQ10G) into the C‐terminal hinge loop allows the protein to form amyloid‐like fibrils, which are shown to contain 3D domain‐swapped molecules (Sambashivan et al., 2005). The authors therefore propose a model of the 3D domain‐swapped fiber (Fig. 13B and C), which contains a run‐away domain swap of the C‐terminal strand and a cross‐b spine formed by the polyglutamine stretches. Like the GNNQQNY cross‐b spine, the modeled polyglutamine spine is made up of two b‐sheets forming a steric zipper. The glutamine side chains interdigitate for a tight packing of the facing sheets (Fig. 13B), and they form hydrogen‐bonded stacks within each sheet (Fig. 13C). Unlike the GNNQQNY spine, the sheets of the RNase A polyglutamine spine are formed by antiparallel stacking of b‐strands; the 3D domain swapping of molecule 1 into 2, 2 into 3, 3 into 4, and so forth, stacks the polyglutamine hinge loops into an antiparallel b‐sheet (Fig. 13C). The remainder of the protein retains its native, enzymatically active structure, and packs around the outside of the cross‐b spine (Fig. 13B). The polyglutamine spine of this model is quite similar to that proposed by Sikorski and Atkins (2005; description to follow), although in the latter, the sheets are untwisted. The 7 strand‐to‐strand twist of the polyglutamine‐ RNase A model recapitulates the helical pitch observed in electron micrographs of the fibrils. We note that although the fibril is modeled as a run‐away domain swap, the data do not rule out fibril formation by direct stacking of domain‐swapped dimers or higher‐order oligomers.

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Fig. 13. 3D domain‐swapping cross‐b spine model of polyglutamine‐RNase A fibrils, from Sambashivan et al. (2005). (A) Ribbon diagram of the crystal structure of bovine RNase A (PDB ID code 1XPS). The monomer’s four disulfide bonds are represented by dotted lines, and N‐ and C‐termini are indicated. The C‐terminal strand, shown in dark gray, is swapped in the fiber. The hinge loop is the site of insertion of the polyglutamine stretch. (B) Surface representation of a section of the 3D domain‐swapped fibril model. The view is down the fibril axis and shows eight molecules. The molecules are colored light or dark to distinguish the two sheets and the swapping. From closest to farthest in the light‐colored sheet, molecule 1 swaps its C‐terminal strand into molecule 2, 2 swaps its strand into 3, and 3 swaps into 4. The polyglutamine stretches of the hinge loops of these molecules form an antiparallel b‐sheet. The same is true for the dark‐colored sheet, with A swapping its C‐terminal strand into B, B into C, and C into D. In the center of the fibril, the glutamine side chains of the two sheets interdigitate, resembling the teeth of a zipper. The globular domains (1–4 and A–D) alternate positions to pack around the cross‐b core. (C) Ribbon representation of six molecules of one sheet of the fibril model. Molecules are colored to distinguish alternately swapped molecules, but they are numbered as in B. This view is perpendicular to the fibril axis, looking at the glutamine side chains (shown as medium‐gray sticks) protruding to form one‐half of the steric zipper. The side chains are stacked, forming hydrogen bonds along the length of the fibril. All panels were generated with Pymol (DeLano, 2002).


IV.

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Models of Natively Disordered Proteins

A third class of models contains those that can be classified as neither Refolding nor Gain‐of‐Interaction models. These are models of proteins whose native state is not structured, or so‐called ‘‘intrinsically disordered’’ proteins (Fig. 2; Dunker et al., 2002). Most of the models fitting the Natively Disordered class describe only segments of proteins. While the full protein may contain regions of identifiable structure in the native state, the fragment responsible for fibril formation does not. Examples include the expanded polyglutamine stretch of huntingtin protein, the C‐terminal segment of the fungal prion HET‐s, the N‐terminal segment of the yeast prion Ure2p, and the amyloid‐b (Ab) polypeptide. The disordered proteins and protein fragments may adopt specific structures on binding a partner molecule, but it is not yet clear whether this induced structure would be relevant to the protein’s fibrillar conformation.

A.

Cylindrical b‐Helices

A cylindrical b‐helix model, called a ‘‘water‐filled nanotube,’’ was proposed by Perutz et al. (2002a) for the fibril structures of several disordered protein fragments, including polyglutamine, a Gln/Asn‐rich peptide of the yeast prion Sup35p, and Ab (Perutz et al., 2002a,b). The model is based on a polyglutamine peptide fibril X‐ray diffraction pattern, in which the authors note the presence of the expected 4.7‐A˚ reflection, an 31‐A˚ reflection, and the absence of the expected 10‐A˚ reflection. They therefore propose a cylindrical, parallel b‐helix with 20 amino acids per coil (Fig. 14A). The authors suggest that the b‐sheet strand‐to‐strand packing accounts for the 4.7‐A˚ reflection, the cylindrical shape explains the lack of an 10‐A˚ sheet‐to‐sheet spacing, and a 20‐residue coil length gives the fibril a diameter of 30 A˚. A prominent 8.3‐A˚ reflection is not interpreted. This b‐helix model is termed a ‘‘water‐filled nanotube’’ (Perutz et al., 2002a) because the large cross section allows an inner pore of 12‐A˚ diameter (Fig. 14B). The parallel stacking of side chains, seen in known b‐helices, is suggested to stabilize the nanotube structure. However, this nanotube model is notably different from known parallel b‐helical structures, as the latter have defined b‐strand and turn regions and little‐to‐no water in their tightly packed cores ( Jenkins and Pickersgill, 2001; Wetzel, 2002). Thus, the stability of the nanotube model has been questioned (Stork et al., 2005; Wetzel, 2002). In a later publication, Kishimoto et al. (2004) proposed the ‘‘water‐filled nanotube’’ as a model for the fibrillar N‐terminal domain of the yeast prion Sup35p. The authors find that hydrated Sup35p fibrils show no 10‐A˚ equatorial reflection in the fiber diffraction pattern, but that dried fibrils

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Fig. 14. Cylindrical b‐helix model of polyglutamine fibrils, from Perutz et al. (2002a). (A) Ribbon diagram of an extended polyglutamine sequence forming a cylindrical, parallel b‐helix. The sequence forms a continuous b‐strand, although the peptide whose diffraction served as the basis for the model would only make up one coil. (B) Surface and stick representation of the model, viewed down the fibril axis. (The top coil is shown as sticks, with the remainder showing van der Waals radii.) The glutamine side chains are proposed to form hydrogen‐bonded stacks parallel to the fibril axis. The large diameter of the cylinder results in a pore down the center. Both panels were generated with Pymol (DeLano, 2002).

do exhibit this reflection. Both dried and hydrated fibrils show the 4.7‐A˚ meridional reflection. Kishimoto et al. (2004) suggest that drying of the fibrils flattens the cylindrical structure into two sheets, creating the spacing that yields the 8‐ to 10‐A˚ equatorial reflection. An alternative interpretation is that in the hydrated fibrils, a weak sheet‐to‐sheet interaction permits several polymorphic packings whose average diffraction intensity is weak. Drying the fibrils produces a stronger, more regular packing, giving rise to the observed 10‐A˚ equatorial reflection. Krishnan and Lindquist (2005) additionally examined Sup35p fibrils using chemical and spectroscopic methods and suggested that part of the protein’s N‐terminal domain forms a protected core, and subunits pack in a head‐to‐head, tail‐to‐tail arrangement. However, these studies (Krishnan and Lindquist, 2005) suggest few further constraints to the conformation of Sup35p in the fibril.

B.

Other Models for Polyglutamine

Critical examination of the ‘‘water‐filled nanotube’’ model has led to suggestions of alternative structures (Sikorski and Atkins, 2005; Stork et al., 2005).


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Sikorski and Atkins (2005) reexamined the polyglutamine peptide fiber diffraction data of Perutz et al. (2002a) and proposed a cross‐b spine model for the fibrillar structure. They found that a sheet‐to‐sheet spacing was indeed suggested by the data, consistent with the previously unexplained 8.3‐A˚ reflection. The model stacks the peptide into flat, untwisted b‐sheets, with glutamine side chains forming hydrogen‐bonded stacks as proposed in the nanotube model (Perutz et al., 2002a, 1994). The side chains from adjacent sheets interdigitate to allow the closer‐than‐expected sheet‐to‐sheet spacing (Sikorski and Atkins, 2005). While parallel and antiparallel b‐sheet models fit the diffraction data equally well, antiparallel sheets were found to be more energetically favorable. Also, the width of certain equatorial reflections led the authors to suggest that the peptide may adopt a b‐hairpin conformation (Sikorski and Atkins, 2005). This model, derived from fiber diffraction data, strongly resembles the model derived from single crystal diffraction of GNNQQNY (Fig. 9; Nelson et al., 2005). Other b‐structured models have been proposed for longer polyglutamine stretches (Bevivino and Loll, 2001; Kajava et al., 2004; Thakur and Wetzel, 2002).

C. HET‐s The prion protein HET‐s is involved in a controlled cell death process in the fungus Podospora anserina (reviewed in Glass et al., 2000; Saupe, 2000). Although the structure of the 289‐amino acid protein has not been determined, CD and NMR spectroscopy suggest that residues 1–227 form a compact, mainly a‐helical domain, and residues 228–289 are flexible and lack a defined secondary structure (Balguerie et al., 2003). CD and FTIR spectroscopy suggest that an increase in antiparallel b‐structure accompanies fibril formation (Dos Reis et al., 2002), and proteolysis experiments reveal a protease‐resistant core spanning residues 218–289 (Balguerie et al., 2003). HET‐s (218–289) also forms fibrils on its own (Balguerie et al., 2003). Ritter et al. (2005) proposed a structural model (Fig. 15) for fibrils of HET‐s(218–289) based on quenched hydrogen exchange NMR, solid‐state NMR, and mutagenesis studies. Each subunit is proposed to form four b‐strands, two of which (b1 and b3) stack parallel in one b‐sheet and face the other two (b2 and b4), which are stacked parallel in the adjacent b‐sheet. The two sheets of this cross‐b spine model are therefore held together by covalent bonds, presumably making it a very stable structure. The structure may also be stabilized by side chain‐stacking interactions, as seen in other parallel b‐structures ( Jenkins and Pickersgill, 2001; Nelson et al., 2005). The HET‐s sequence spanning b1 to b2 shares 28% identity

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Fig. 15. Cartoon representation of the cross‐b spine model of HET‐s fibrils, based on Fig. 2 of Ritter et al. (2005). The stretch of residues 218–289 is shown as the four central b‐strands, b1–b4, plus the connecting loops. N‐ and C‐termini are indicated.

with that of b3 to b4 (Ritter et al., 2005), so in many cases the b1–b3 stack (or b2–b4 stack) could have identical side chains stacked one on the other.

D. Ure2p The prion protein Ure2p is involved in regulating nitrogen catabolism in yeast (reviewed in Komar et al., 1999). Soluble Ure2p exists as a native dimer, in which residues 1 to 95 form a flexible, largely unstructured segment, and residues 95–354 make up a mainly a‐helical, globular domain with a GST‐like fold (Fig. 16A; Bousset et al., 2001; Umland et al., 2001). Different research groups have examined Ure2p fibrils having different properties. In the following paragraphs, we describe models of amyloid‐like and non‐ amyloid‐like Ure2p fibrils. For a more in‐depth discussion, we refer the reader to the chapter in this volume by Baxa et al. (2006). Certain filaments of Ure2p give a cross‐b diffraction pattern (Baxa et al., 2005), and scanning transmission electron microscopy (STEM) images reveal the filaments to be composed of a 4‐nm‐diameter core surrounded by globular appendages (Baxa et al., 2003). Protease treatment of these filaments produces a 4‐nm‐diameter filament composed of Ure2p fragments roughly spanning residues 1–70 (Baxa et al., 2003). Likewise, the Ure2p segment 1–65 forms b‐sheet rich, amyloid‐like filaments in vitro (Taylor et al., 1999) and induces the prion phenotype in vivo (Masison and Wickner, 1995).


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Fig. 16. Parallel superpleated b‐structure proposed for Ure2p amyloid‐like fibrils, from Kajava et al. (2004). (A) Ribbon diagram of a dimer of Saccharomyces cerevisiae Ure2p C‐terminal domains (PDB ID code 1G6W), generated with Pymol (DeLano, 2002). The monomers are colored in light and dark gray and are viewed down the twofold symmetry axis. The N‐ and C‐termini are indicated, and residue 137 is denoted by an open arrow. (B) Cartoon representation of the parallel superpleated b‐structure proposed for the N‐terminal 70 residues of Ure2p in the fibril. This view down the fibril axis shows the stacking of N‐termini with a twist of 3 . For reference, the calculated twist based on fibril pitch ranges from 0.7 to 3.4 from one monomer to the next (Kajava et al., 2004). (C) Cross section of the Ure2p amyloid‐like fibril model, showing the parallel superpleated b‐structure at the N‐terminus, and various positions possibly occupied by the globular C‐terminus (gray ovals). Panels B and C are based on Fig. 4 of Kajava et al. (2004).

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Based on these and related data, Kajava et al. (2004, 2005) have proposed the ‘‘parallel superpleated b‐structure’’ as a model for Ure2p filaments and other amyloid‐like fibrils. In this model, the N‐terminal 70 residues of the Ure2p monomer adopt a serpentine conformation, zigzagging to form a set of b‐strands that interact via their side chains (Fig. 16B). Each strand is four residues long and strands are separated by three‐ residue turns. The serpentine N‐terminal segments stack parallel and in‐register in the direction of the fiber axis, forming a cross‐b spine of parallel b‐sheets. The C‐terminal globular domains pack around the sides of the spine (Fig. 16C). The parallel, in‐register, stacks allow stacking of identical side chains for added stability. However, the elongated shape of this spine (27 nm) (Kajava et al., 2004) does not match the core size measured by STEM (4 nm) for the intact or protease‐treated Ure2p filaments (Baxa et al., 2003). Solid‐state NMR studies of fibrillar Ure2p10–39 (Chan et al., 2005) suggest that this peptide also packs in a parallel, in‐register b‐structure, though the regions of sequence comprising the strands versus the turns are suggested to differ from those of the Kajava model (2004). Fay et al. (2005) have proposed a completely different model for Ure2p fibril structure. Their model is based on data which suggest that Ure2p fibrils do not have a cross‐b structure (Bousset et al., 2003) and that the C‐terminal globular domain is tightly involved in the fibrillar scaffold (Bousset et al., 2004). In this model (Fay et al., 2005), portions of the N‐ and C‐terminal regions, specifically residues 6 and 137, are in close proximity in the fibril, and the C‐terminal domain retains a native‐like structure. There is evidence that this non‐amyloid‐like fibril can convert to the cross‐b‐containing filament with heat treatment, incubation at low pH (Bousset et al., 2003), or extensive drying (Fay et al., 2005), but it is unclear what sort of structural change might link the two fibril types.

E. Amyloid b The Ab polypeptide may be the most widely studied of all amyloid‐ forming sequences because it forms amyloid deposits in the brains of patients with Alzheimer’s disease (Selkoe, 1994, 1996). Ab is a 39‐ to 43‐amino acid cleavage product of the Ab precursor protein (Glenner and Wong, 1984; Kang et al., 1987; Masters et al., 1985; Prelli et al., 1988; Vassar et al., 1999). Ex vivo fibrils of Ab show X‐ray diffraction reflections at 4.76 and 10–11 A˚ (Kirschner et al., 1986), and fibrils formed in vitro give a cross‐b diffraction pattern (Inouye et al., 1993; Malinchik et al., 1998). In addition to studies of full‐length Ab, there have been a myriad of studies examining the fibril‐ forming and structural properties of shorter segments of Ab (Antzutkin


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et al., 2002; Balbach et al., 2000; Benzinger et al., 1998, 2000; Bond et al., 2003; Burdick et al., 1992; Burkoth et al., 2000; Castano et al., 1986; D’Ursi et al., 2004; Fraser et al., 1991, 1994; Gordon et al., 2004; Gorevic et al., 1987; Gregory et al., 1998; Halverson et al., 1990; Hilbich et al., 1991; Inouye et al., 1993; Ippel et al., 2002; Kellermayer et al., 2005; Kirschner et al., 1987; Lansbury et al., 1995; Liu et al., 2004; Petkova et al., 2004; Petty and Decatur, 2005; Serpell and Smith, 2000; Serpell et al., 2000a; Shanmugam and Jayakumar, 2004; Sikorski et al., 2003; Tjernberg et al., 1999; Wood et al., 1995). Taken together, these examinations reveal that peptide sequence, fragment length, and fibrillization conditions (e.g., pH) all affect the structure of the fibril; fibrils may be composed of parallel or antiparallel sheets, peptides may be extended and/or contain turns, and multiple registers of the stacked peptides are possible. For brevity, we focus on recent structural models of full‐length Ab peptides. Petkova et al. (2002) proposed a fibril model for Ab1–40 based on constraints from solid‐state NMR (ssNMR) experiments. The ssNMR data suggest that residues 1–10 are disordered, residues 9–21 and 30–36 consistently adopt b‐strand conformations, and residues 23, 24, 25, and 29 may adopt non‐b‐strand conformations. In addition, prior ssNMR (Antzutkin et al., 2000, 2002; Balbach et al., 2002) and site‐directed spin‐labeling (To¨ro¨k et al., 2002) experiments suggested that full‐length Ab peptides stack parallel and in‐register. The authors (Petkova et al., 2002) therefore propose a parallel‐stacked hairpin‐like structure (Fig. 17A and B). Residues 12–24 and 30–40 make up the 2 b‐strands, and residues 25–29 form a bend in the chain, bringing the side chains of the 2 strands into proximity (Fig. 17A). The hairpins stack parallel and in‐register along the length of the fibril, forming a cross‐b spine. Lateral packing of hairpins is driven by hydrophobic interactions between C‐terminal strands; two stacks of hairpins pack to form a protofilament (Fig. 17B). This protofilament cross section is supported by mass‐per‐unit‐length measurements, suggesting two peptides are contained within each 4.8‐A˚ ‘‘layer’’ of the protofilament, and by EM measurements of the smallest fibril diameters: 5010 A˚. The authors also model an association mechanism for producing fibrils with larger diameters (Fig. 17B). A related fibril model for Ab1–40 was proposed based on scanning proline mutagenesis (Williams et al., 2004) and molecular modeling (Guo et al., 2004). This model proposes that residues 15–21, 24–28, and 31–36 form 3 b‐strands, with 2 intervening turns formed by residues 22–23 and 29–30 (Fig. 17C). Residues 17 and 34 are placed in close proximity, as double cysteine mutants at these positions form disulfide bonds on oxidation after fibrillization (Shivaprasad and Wetzel, 2004). Since fibrils with this triangular cross section would not be expected to show an 10‐A˚

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Fig. 17. Models of Ab(1–40) fibrils, viewed down the fibril axis. (A, B) Cartoon representations of the hairpin model proposed by Petkova et al. (2002). (A) The proposed b‐strands span residues 9–24 and 30–40, with a main‐chain bend spanning residues 25–29. Aspartate 23 and lysine 28 are proposed to form a salt bridge (dotted line) based on distance constraints provided by ssNMR. The hairpins stack in‐register to form two parallel b‐sheets. (B) Two hairpin stacks pack together to form the smallest observed fibrils, or protofilaments, burying the hydrophobic residues of the C‐terminal strand. Two or more protofilaments may pack together to form thicker fibrils. (C) Cartoon representation of the parallel b‐helix‐like model proposed by J. T. Guo and Y. Xu (unpublished; model shown in Fig. 1 of Shivaprasad and Wetzel, 2004). The gray oval highlights residues 17 and 34, proposed to sit in close proximity.

equatorial reflection, Guo et al. (2004) propose a 10‐A˚ sheet‐to‐sheet packing between triangular stacks.

V.

Fibril Properties and Their Relation to Structural Models

Amyloid, in its strictest definition, is an extracellular deposit, composed of protein fibrils, proteoglycans, glycosaminoglycans, the amyloid P component, and sometimes other plasma proteins (Pepys, 2006, and references therein; Westermark et al., 2005). The component fibrils display an elongated, unbranched morphology; bind Congo red to give a characteristic green birefringence; and give a cross‐b X‐ray diffraction pattern. Many other protein fibrils share some, but not all, of these characteristics, and are not strictly classed as ‘‘amyloid.’’ They are, however, important for the

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Table I Consistencies of the Three Model Classes with the Common Properties of Amyloid‐like Fibrilsa Gain‐of‐Interaction models Common fibril property Fibrillar morphology Protofilament substructure Variable morphology Cross‐b diffraction Increase in b‐structure Congo red binding with birefringence Self‐only association Cooperative kinetics of formation Extreme stability

Refolding models

Natively Disordered models

I

II

III

IV

ü ü

ü ü

ü ü

ü ü

ü? ü

? ?

? ü

? ?

? ü

?

ü

?

ü

ü ü X b‐helices ü

?

?

?

?

?

? ?

ü X

ü ?

ü ?

ü ?

? ?

?

ü

ü

ü

ü

?

? ü ü ü X b‐helices ü a‐helical and mixed proteins ? all‐b proteins ?

a

Key: ü, consistent; X, inconsistent; ?, unclear. Properties shown in bold are defining properties of amyloid fibrils.

physicochemical examination of the common underlying structure of amyloid fibrils, and they have revealed other characteristics of this broader group of ‘‘amyloid‐like’’ fibrils. The Refolding, Gain‐of‐Interaction, and Natively Disordered classes of fibril models are at least partly consistent with the common properties of amyloid and amyloid‐like fibrils. We summarize consistencies, inconsistencies, and uncertainties linking model class and amyloid property in Table I. In the following paragraphs, we describe these properties and discuss the extent to which they may be explained by the various classes of models.

A.

Fibrillar Morphology

The common elongated, unbranching morphology of amyloid‐like fibrils suggests an ordered arrangement of subunits, where inter‐subunit interactions are strongest along the long dimension of the fibril. While the fibrillar morphology goes hand‐in‐hand with the cross‐b structure of the

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fibrils (discussed in Section V.D), there are many examples of fibrillar protein structures composed of other arrangements of secondary structure (e.g., collagen, keratin, actin, tubulin, and silk). It has been suggested that the amyloid conformation represents a default or generic conformation for a polypeptide chain (Dobson, 1999). In this case, the Refolding models would be consistent with the amyloid fibrillar morphology. However, one might expect that the extensive conformational changes required by the Refolding models would expose suitable interaction surfaces in more than one dimension, resulting in disordered or alternately ordered aggregates. The Refolding models do not clarify how proteins make ordered, linear, intermolecular associations. The Gain‐of‐Interaction models seem more consistent with the amyloid fibrillar morphology; inter‐subunit interactions are only formed through a newly exposed segment, while the majority of the protein retains its native structure and dissuades further interactions. In the direct‐stacking models (Type I), a protein subunit requires two interaction surfaces to form a fibrillar structure. We note that it may not be necessary for a protein to expose two new interaction surfaces; in the models of TTR and SOD in preceding sections, each subunit makes use of a preexisting dimeric interface, such that the gain of just one new interface is sufficient for fibril growth. In the cross‐b spine models (Types II and IV), the cross‐b structure promotes growth in one dimension (discussed in Section V.D). In the 3D domain‐swapping models (Types III and IV), a run‐away domain swap naturally assumes a linear arrangement.

B.

Protofilament Substructure

Many amyloid fibrils seem to be made up of smaller protofilaments. Although the number of protofilaments per fibril varies, the protofilaments have a fairly consistent diameter of 30 A˚ (Serpell et al., 2000b; Shirahama and Cohen, 1967; Shirahama et al., 1973). For some proteins, for example TTR, the protofilament diameter matches that of the native protein, suggesting that a Gain‐of‐Interaction model is plausible (Serpell et al., 1995). For other proteins, for example the SH3 domain, the protofilament seems too small to accommodate the native protein structure, suggesting that a Refolding model is plausible ( Jimeńez et al., 1999). We note, however, that measurements of protofilaments are not exact. Protofilament diameters are often measured from negative‐stain electron micrographs, which may introduce errors and/or variability in size due to drying‐induced deformation of the sample, sample‐stain interactions, and variable staining (Chapman et al., 1990; Kiselev et al., 1990). Additionally,


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cryo‐EM suffers from small scaling uncertainties (as an example, see Chiu et al., 1998), and exact edges are difficult to define due to the low contrast between protein and ice (Saibil, 2000). Thus, the protofilament dimensions and morphology determined by EM give only a rough idea of the conformation of the component protein.

C.

Variable Morphology

It is common among fibril‐forming proteins that a single protein can form fibrils of varied morphology. Variations may arise at the level of subunit conformation, which seems to be the case for the prion proteins, or at the level of protofilament packing, as described in the preceding for insulin. Refolding and Natively Disordered models seem consistent with both types of variable morphology. A protein that has unfolded extensively or is normally disordered might be able to fold into several different, but related, cross‐b structures. This is especially true in Refolding models, since backbone interactions are suggested to drive the folding process. If the protein does adopt only one fold at the subunit level, the resulting protofilaments may still have several favorable interaction surfaces, allowing for alternate protofilament packings. The Gain‐of‐Interaction models differ from the Refolding models in that a limited and specific region of the protein is responsible for fibril formation. It therefore seems less likely that these subunits would adopt alternate conformations in forming protofilaments. However, certain cross‐b spines might be able to adopt multiple registries of the intersheet interdigitation, thus allowing polymorphic packing within the protofilament. Also, it seems reasonable that gained‐interaction protofilaments could pack together in multiple ways, as the nature of inter‐protofilament interactions is still unclear.

D. Cross‐b Diffraction Pattern The cross‐b diffraction pattern is a defining feature of amyloid fibrils, so most fibril models incorporate the cross‐b structure. There are, however, examples of models in each of the three classes that do not fit with this pattern. For example, the b‐helical models of PrP (Refolding; Govaerts et al., 2004), polyglutamine (Natively Disordered; Perutz et al., 2002a), and Sup35p (Natively Disordered; Kishimoto et al., 2004) would not be expected to give a cross‐b diffraction pattern; they would show the 4.7‐A˚ meridional reflection, but not the defining 10‐A˚ equatorial reflection. We note that Kishimoto et al. (2004) argue that the 10‐A˚ reflection is an artifact of

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drying, and so they suggest that models lacking this sheet‐to‐sheet spacing are more accurate. In the Gain‐of‐Interaction models, nothing about Types I and III (the direct‐stacking and 3D domain‐swapping models) requires that they give a cross‐b diffraction pattern, or even that they contain b‐structure. In these cases, the proteins would need to have native b‐structure and pack in a way that gives the cross‐b pattern. The cross‐b structure explains the fibrillar morphology of amyloid and amyloid‐like fibrils. The strongest noncovalent interactions in the cross‐b structure are the backbone hydrogen bonds. These interactions dominate the growth of the fibril, and hence are parallel to the long fibril axis. Side chain interactions govern sheet‐to‐sheet packing, thus controlling lateral growth of the fibril or protofilament. Fibril growth in the lateral dimension is restricted and is slower than growth in fibril length because the electrostatic properties and sizes of the side chains must be complementary for sheets to pack together, and because, on average, side chain interactions are weaker than backbone hydrogen bonds.

E. Increase in b‐Structure Several proteins seem to undergo an increase in b‐structure in converting from the native to fibrillar state. Dramatic increases in b‐structure, for example as reported for insulin and myoglobin, lend support to the Refolding models, especially because these native proteins are largely a‐helical. Slight changes in secondary structure, on the other hand, lend support to the Gain‐of‐Interaction models, in which much of the protein retains its native structure. In all cases, it is important to weigh the evidence for structural change while considering the experimental source. CD and FTIR spectroscopy are techniques commonly used to measure the secondary structures of the native versus amyloid‐like fibrillar states. While both techniques can be useful in estimating secondary structure, each is limited in accuracy by the sample, set up, and methods of spectral deconvolution and interpretation. In CD spectroscopy, for example, high protein concentrations can lead to aggregation and light scattering, increasing the noise of the measurement. While measurement from thin films is an alternative for studying bulky samples like amyloid fibrils, it is difficult to define the essential values of protein concentration and path length with this experimental setup (McPhie, 2004). CD and FTIR spectroscopies each have certain solvent requirements. In CD spectroscopy, buffer and solvent solutions must be carefully selected so as to avoid those that absorb in the UV or are optically active (Pelton and McLean, 2000). In FTIR spectroscopy, water absorbs strongly in the amide I


269

region—a key region for estimating secondary structure. Aqueous samples must therefore (1) be dried, which can alter their secondary structure; (2) be dissolved in D2O, which may skew secondary structure determination if the isotope exchange is incomplete; or (3) be highly concentrated such that the contribution from water is small enough to be subtracted ( Jackson and Mantsch, 1995). Both methods are also limited in accuracy of secondary structure determinations because spectral peaks must be deconvolved; estimates are made of the overlapping contributions of different structural regions. These estimates may introduce error based on the reference spectra used and because deconvolution methods equate crystallographic secondary structure with the secondary structure of the protein in solution (Pelton and McLean, 2000). As amyloid fibrils are neither crystalline nor soluble, there may be even greater error in estimates of secondary structure. To compound the problem, estimates of b‐sheet content are less reliable than those of a‐helix, because of the flexibility and variable twist of b‐structure (Pelton and McLean, 2000). In addition, b‐sheet and turn bands overlap in FTIR spectroscopy ( Jackson and Mantsch, 1995; Pelton and McLean, 2000). Side chains also contribute to spectral peaks in both methods, and they can skew estimates of secondary structure if not properly accounted for. In FTIR spectra, up to 10–15% of the amide I band may arise from side chain contributions ( Jackson and Mantsch, 1995). These sources of error in estimating secondary structure content from CD and FTIR spectra suggest that the estimates, especially for amyloid fibrils, should be considered qualitative.

F. Congo Red Binding and Birefringence The birefringence of amyloid‐bound Congo red indicates an ordering of the dye, and therefore an ordered arrangement of proteins in the fibril. Although there have been several proposals as to the mechanism of Congo red binding to amyloid fibrils (Carter and Chou, 1998; Gueft and Ghidoni, 1963; Inouye and Kirschner, 2005; Kajava et al., 2004; Klunk et al., 1989; Roterman et al., 2001; Turnell and Finch, 1992; Wolman and Bubis, 1965), the specific structural feature(s) recognized by Congo red remains unclear. It is therefore difficult to say whether any of the classes of fibril models are consistent with Congo red binding and birefringence.

G. Self‐Association Each type of amyloid fibril is formed from one type of protein (Westermark et al., 2002), suggesting a specificity in fibril formation. This specific

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self‐association may result from sequence specificity, consistent with the Gain‐of‐Interaction models, as they propose that the protein provides a specific interaction surface. Self‐association may also be a result of destabilization specificity, in that the conditions in vivo causing destabilization of the protein are specific to that one protein. This destabilization specificity would seem to better explain the Refolding models, as their formation is proposed to be governed by non‐sequence‐specific backbone interactions (Fa¨ndrich et al., 2001). However, unlike nonspecific aggregates, amyloid fibrils are linear, ordered structures. Therefore, while there may be a destabilization specificity in vivo, it seems likely that sequence‐specific interactions promote ordered aggregation.

H. Cooperative Kinetics of Formation In vitro, fibril formation by several proteins displays an initial lag phase, followed by a rapid increase in aggregation (reviewed in Rochet and Lansbury, 2000). Introduction of fibrillar seeds eliminates the lag phase. These cooperative aggregation kinetics suggest that fibril formation begins with the formation of a nucleus and proceeds by fibril extension. The structure of the nucleus must therefore act as a template for the protein’s conformation in the fibril. As the structural requirements for templating are unclear, it is difficult to assess the consistency of the model classes with this feature of fibril formation. We have described one possible templating mechanism for the cross‐b spine of GNNQQNY (Nelson et al., 2005). One model type that is especially difficult to explain in terms of cooperative aggregation kinetics is the direct‐stacking model (Type I, Gain‐ of‐Interaction). Direct‐stacking models have two, seemingly independent, interaction surfaces. How does binding of one surface to the growing amyloid fibril create a template for binding at the other surface? Perhaps it does not. Hurshman et al. (2004) suggest that TTR fibrillization in vitro does not follow nucleation‐dependent kinetics; after the rate‐limiting step of dissolution of the TTR tetramer into a monomer, fibril formation by monomeric TTR may be a downhill polymerization process. In addition, Gosal et al. (2005) have shown that b2m follows two competing pathways to form distinct types of amyloid‐like fibrils in vitro: one pathway shows nucleation‐limited aggregation kinetics, and the other shows non‐nucleation‐ dependent kinetics. Perhaps, then, fibril formation by direct stacking is a non‐nucleation‐dependent, noncooperative process, and is an alternate route to amyloid fibril formation.


I.

271

Extreme Stability

The extreme stability of amyloid and amyloid‐like fibrils is difficult to understand in terms of the three classes of fibril models. For the Refolding models, it has been suggested that the amyloid conformation is a default conformation for a polypeptide chain (Dobson, 1999). However, these models do not give a clear indication of what types of interactions differ in the amyloid conformation versus the native conformation, and so it is unclear why the amyloid conformation should be more stable. Also, it seems that the elevated protein concentrations associated with fibril formation might disproportionately favor nonspecific aggregation of the destabilized intermediate over amyloid fibril formation. In the Gain‐of‐Interaction models, new interactions in only a limited region of the protein must account for the gained stability. We have considered the energetics of cross‐b spine formation by GNNQQNY (Nelson et al., 2005), and we conclude that peptide concentration is the key determinant of stability of the peptide fibrils. At low monomer concentration, not enough intermolecular interactions are formed to balance the peptide’s loss of entropy in polymerization. At high monomer concentration, the likelihood of fibril‐inducing interactions increases, and the cooperativity of formation produces a fibril with lower free energy than that of the monomer. The cooperativity in the formation of 3D domain‐ swapped fibrils would similarly be expected to stabilize the fibril at high protein concentration. Additional factors may influence fibril stability in vivo. The glycan molecules and amyloid P component found in amyloid deposits may stabilize amyloid fibrils (Pepys, 2006, and references therein). Also, monomer concentration is important to amyloid formation and stability in vivo; several amyloidoses are associated with elevated levels of the fibril precursor protein, and deposits regress when the levels of the precursor protein are sufficiently reduced (Pepys, 2006, and references therein).

VI.

Conclusions

In this chapter, we present several examples of structural models for amyloid fibrils, which we group into general classes. None of these general model classes can completely explain the common properties of amyloid and amyloid‐like fibrils; however, the Gain‐of‐Interaction models with a cross‐b spine seem most consistent with what is known. These models combine the structural aspect of the cross‐b spine with the specificity of sequence‐dependent interactions to explain the observed diffraction, stability, and self‐only association of amyloid fibrils. It is also possible

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that a variety of fibril structures exists, such that more than one type of model will be needed to represent the diversity of fibrils. The structural bases for the common fibril properties should become clearer as models are updated and techniques for examining the atomic‐level structural details of fibrils are improved. These structural details are key to understanding how amyloid fibrils form and to developing therapeutics for certain amyloid diseases.

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AUTHOR INDEX

A Abraham, C., 14, 277 Abraham, D. J., 277 Abrahams, J. P., 123 Abrahamson, M., 272, 275–276 Abrescia, N. G., 99, 118–119 Adams, J. M., 91 Adams, M. J., 272 Adams, W. W., 49 Adrian, M., 119, 232 Aebi, M., 176 Aebi, U., 12, 14, 175–176, 229–233, 277 Agard, D. A., 213, 282 Aggeli, A., 112, 118 Aguet, M., 173, 206 Aguzzi, A., 135, 172, 178 Ahrazem, O., 14 Akai, H., 22, 25, 28, 36, 38, 42 Akita, M., 62, 67, 90 Alber, B. E., 93 Albert, A., 14 Albiges-Rizo, C., 122 Almo, S. C., 92 Alperovitch, A., 213 Altenbach, C., 280 Altman, G. H., 31, 42 Amadei, A., 207 Amarante, P., 282 Amoresano, A., 278 Amorin, M., 206 Andersen, S. M., 94 Anderson, D. E., 92 Anderson, J. P., 31, 42 Anderson, R. E., 92 Anderson, V. E., 232 Andersson, R., 94 Andrieu, J. P., 121 Andronesi, O. C., 231 Anfinsen, C. B., 37, 39, 42, 245, 272

Angeretti, N., 207, 213 Antholine, W. E., 206 Antony, T., 231 Antonysamy, S., 91 Antonyuk, S., 275 Antzutkin, O. N., 13–14, 85, 91, 156, 172, 229, 233, 272, 279 Apetri, M. M., 232 Appel, R. D., 92 Aravinda, S., 206 Arcdiacono, S., 49 Argos, P., 208 Arisaka, F., 47, 93, 120–121 Arispe, N., 229 Armenante, M. R., 274 Arnaudov, L. N., 39, 42 Arnberg, N., 118 Arnold, E., 85, 95 Arnott, S., 206 Arnsdorf, M. F., 178 Aronoff-Spencer, E., 92, 206 Artzner, F., 213 Arutzen, C. J., 42 Arvinte, T., 175, 230 Arzt, S., 91 Asakura, S., 211 Asakura, T., 25–26, 28, 30–31, 38, 42–43, 47, 49–50, 53 Asano, S., 233 Ashburn, T. T., 14, 46, 120, 278 Ashida, J., 43 Ashman, K., 123 Astbury, W. T., 3, 13, 272 Atkins, E. D. T., 45, 48–49, 119, 211, 213, 278, 281 Attinger, A., 209, 276 Auger, M., 14, 120, 278 Autenried, P., 206 Avdievich, N. I., 206 Avtges, P., 52

283

284

AUTHOR INDEX

Awan, T., 207, 213 Awni, L. A., 277 Azimova, R., 233 Azuma, M., 36, 43

B Babu-Khan, S., 282 Bachinger, H. P., 119 Badger, J., 63, 68, 70, 75, 86–87, 91 Bagchi, S. N., 208 Bai, M., 148, 172 Baier, M., 213 Bailey, K., 272 Bain, K., 91 Bairoch, A., 92 Baker, E. N., 272 Balaram, P., 206 Balbach, J. J., 10, 13–14, 91, 172, 229, 233, 272, 275, 279 Balbirnie, M., 14, 121, 174, 177, 206–207, 272, 278 Balch, W. E., 231 Baldus, M., 231 Baldwin, M. A., 175, 177, 178, 180, 207, 209, 211, 213, 275, 279, 282 Baldwin, R. L., 27, 43 Balguerie, A., 134, 137, 140, 147, 172–173, 272 Ball, H. L., 209 Ballicora, M. A., 93 Bamford, D. H., 118–119, 121, 123 Bamford, J. K., 118–119 Banerjee, A., 211 Bann, J., 119 Baranov, V., 279 Barford, D., 59, 61, 92 Barge, A., 121–122 Bark, N., 213 Barlow, A. K., 232 Barnham, K. J., 210 Barnikol, H. U., 277 Baron, G. S., 208 Baroni, F., 230 Barron, L. D., 211 Barrow, C. J., 210 Barry, M. A., 121 Bartlam, M., 233, 281 Barton, E. S., 102, 118

Baskakov, I. V., 10, 13, 129, 134, 173, 176, 210, 215 Bastiaansen, C. W. M., 48 Bastidas, R. B., 178, 212, 279 Bateman, A., 82, 91 Bathany, K., 172, 272 Batiyenko, Y., 91 Bauer, H. H., 229 Baum, J., 113, 118 Baumann, U., 56–57, 63, 68, 70, 75, 85, 91–92 Bax, A., 178 Baxa, U., 10, 14, 89, 93, 95, 120, 122–123, 125, 139–141, 144–149, 151, 155, 157, 162, 164, 173, 176–178, 180, 206, 210, 272, 277, 282 Baxter, H. C., 208 Bayer, P., 213 Bayley, P. M., 208 Beaman, T. W., 64, 67, 69, 87, 91 Beasley, J. R., 123 Beaulieu, L., 43, 50 Becker, M. A., 47, 51 Becker, O. M., 210 Becker, S., 231 Beckmann, J. S., 178 Bedzyk, L. A., 45 Beek, J. D., 18, 21, 30, 43 Begueret, J., 174–175, 274 Beighton, E., 45, 119 Bell, M., 118 Bello, J., 277 Bellotti, V., 278 Belousova, N., 120 Belrhali, H., 121, 173, 273 Benavente, J., 119 Bendheim, P. E., 212, 280 Benedek, G. B., 48, 233 Benen, J. A., 95–96 Benevides, J. M., 119 Bennett, B. D., 282 Bennett, M. J., 272, 280 Bennhold, H., 229 Benson, M. D., 180, 273, 281–282 Benson, S. D., 123 Bentley, J. D., 92 Benyamini, H., 273 Benzinger, T. L. S., 10, 13, 156, 173, 230, 273–274, 276 Bergelson, J. M., 100, 118, 122

AUTHOR INDEX

Berger, B., 91 Bergmann, M., 213 Bergseid, M. G., 91 Bergstrom, A. L., 206 Berman, H. M., 61, 91 Bernard, A. R., 91 Bernhagen, J., 213 Bernstein, D. T., 52 Berriman, J. A., 122, 175, 178, 212, 279 Bertani, I., 207 Berthet-Colominas, C., 119 Bertoldi, M., 275 Bertoluzza, A., 49 Bevivino, A. E., 273 Bewley, M. C., 99–101, 118 Beyreuther, K., 213, 276–278 Bhat, K. S., 13, 174, 274 Bhat, T. N., 91 Bhatia, R., 232 Billeter, M., 180, 212, 214, 282 Binda, C., 58, 63, 68, 79, 87, 91 Bini, E., 21, 23, 25, 44–45 Birkett, C., 206 Bjorkman, P. J., 280 Black, G. W., 95, 122 Blackledge, T. A., 46 Blackley, H. K., 230 Blake, C. C. F., 3, 13, 37, 50, 115, 123, 215, 233, 273, 281 Blanch, E. W., 211 Blanchard, J. S., 91–92 Blondelle, S. E., 213 Bloom, J. D., 178 Bluethmann, H., 173 Blume, A., 213 Blundell, T. L., 272 Bocharova, O. V., 10, 13, 134, 173 Bode, W., 273 Boden, N., 118 Bohrmann, B., 232 Bolognesi, M., 233 Bolton, D. C., 212, 280 Bond, J. E., 120, 206, 209, 273 Bonjour, S., 208 Bonneu, M., 177 Bonthrone, K. M., 39, 44 Booker, G. W., 273 Bossers, A., 212 Bossi, R. T., 91, 96 Botto, R. E., 13, 173, 230, 273–274, 276

285

Bouchard, M., 176, 273, 277 Boulanger, P. A., 113, 121 Bourne, P. E., 91 Bourne, Y., 95 Bousset, L., 136, 143, 146, 148–149, 151, 173, 175, 179, 273–275 Bouzamondo-Bernstein, E., 177 Bowman, K. A., 212, 280 Boyd, J., 273 Brachmann, A., 141–143, 167–169, 172–173, 180 Brack, A., 49 Bradbury, E. M., 3–4, 13 Bradford, C., 15, 180 Bradley, M. E., 174 Bradley, P., 76, 89, 91 Bram, A., 30, 44 Branden, C. I., 44, 50 Branham, C., 47 Braun, F. N., 39, 44 Brennan, M. J., 94 Breydo, L., 173 Briki, F., 173, 273 Brodsky, B., 113, 118 Bromley, E. H., 232 Brookes, V. L., 50, 53 Brooks, E. G., 92 Broome, B. M., 36, 44 Brough, D. E., 122 Brown, C. J., 175, 274 Brown, L., 13, 206 Bruce, M. E., 166, 173, 206 Brunner, H., 213 Bruschi, M., 213 Bryant, P. K. III, 212 Brzin, J., 273 Buard, J. J., 89–89, 91, 282 Bucciantini, M., 230, 233 Buchanan, M. D., 91 Buchanan, S. G., 91 Bucher, P., 76, 91 Buchner, J., 179 Buder, K., 275 Budisa, N., 123 Bueler, H., 132, 173, 206 Bugiani, O., 207–208, 212–213 Buntkowsky, G., 279 Burda, M. R., 96, 108, 114, 118, 123 Burdick, D., 273 Burke, M. J., 273

286

AUTHOR INDEX

Burkhard, P., 87, 91 Burkoth, T. S., 13, 173, 230, 273–274, 276 Burley, S. K., 206 Burlingame, A. L., 213 Burmeister, W. P., 99, 118–119 Burnett, R. M., 118, 123 Burns, C. S., 206 Burton, D. R., 177–178, 210, 212, 279 Butcher, S. J., 118, 123 Butler, D. A., 178 Buxbaum, J. N., 180, 282 Byeon, I. J., 177

C Cahn, R. W., 45 Calabro, T., 42 Calder, L. J., 123 Calkins, E., 230, 274 Callaway, D. J., 281 Calvo, P., 213 Calzolai, L., 180, 206, 211, 214, 282 Cambillau, C., 122 Campbell, I. D., 273, 276 Campbell, J. A., 121 Canale, C., 233 Canciani, B., 208, 213 Canetti, M., 31, 34, 44 Cannon, M. J., 40, 44 Cao, L., 207 Capobianco, R., 213 Cappello, J., 42 Caramelli, M., 207 Cardoso, I., 230 Carey, P. R., 232 Carp, R. I., 166, 173 Carrington, E., 46 Carter, D. B., 274 Carter, S. A., 232 Cary, R. B., 274 Case, S. T., 52 Casem, M. L., 36, 44 Cashikar, A. G., 178 Caspar, D. L. D., 174–175, 207 Caspi, S., 207 Cassese, T., 282 Castano, E. M., 274–275, 280 Castedo, L., 206

Castilla, J., 134, 174 Caughey, B. W., 13, 174, 179, 207–208, 210, 212–213, 274 Caughey, W. S., 3, 13, 150, 174, 207, 274 Cayabyab, A., 274 Cerritelli, M. E., 72, 91, 105, 108, 118, 121 Cesari, F. C., 30, 48 Chabry, J., 206–207 Chaignepain, S., 172–173, 272 Chan, J. C., 10, 13, 156, 159, 174, 274 Chandler, H. D., 46 Chantalat, L., 94 Chao, K. L., 92 Chapman, J. A., 274 Chappell, J. D., 98, 100–101, 103, 106, 118, 121 Charbit, A., 123 Charles, I. G., 92, 119 Charnock, S. J., 95, 122 Chase, S., 51 Chen, B.-L., 111, 118 Chen, D. J., 274 Chen, J. S., 13, 42, 174 Chen, M., 213 Chen, X., 31, 36–37, 44, 48, 52–53, 213 Chen, Y., 213 Cheng, N., 93, 121, 172–173, 179, 206, 272, 281 Cherif-Cheikh, R., 213 Chernoff, Y. O., 174 Cherny, D., 231 Cherny, I., 116, 119 Cherny, R. A., 210 Chesebro, B., 207, 210 Chien, P., 146, 148, 167, 174, 179 Chiesa, R., 207 Chipman, P. R., 93, 120 Chiti, F., 37, 44, 144, 174, 230, 233 Chiu, C. Y., 274 Chiu, T. K., 173, 206, 272 Cho, H. S., 60, 91 Chong, A., 208 Chothia, C., 64, 91, 230 Chou, K. C., 274 Chou, P. Y., 74, 91 Chouin, E., 121 Chree, A., 206 Christen, B., 211 Christopher, J. A., 91 Chroboczek, J., 120–121, 123–124

287

AUTHOR INDEX

Chromy, B. A., 232 Chung, D. S., 48 Chunyu, L., 48 Claborn, K., 47 Clantin, B., 62, 67, 70, 83–85, 88, 91 Clark, A., 44, 174, 231, 278 Clarke, A. R., 209 Clayton, D. F., 230 Cleasby, A., 73, 91 Cleaver, R., 93 Cockburn, J. J., 99, 118–119 Coda, A., 91, 96 Cohen, A. S., 180, 213, 230, 233, 274, 281–282 Cohen, C., 82, 91 Cohen, F. E., 14, 119, 173, 175–178, 180, 207–213, 276, 279, 282 Colacino, S., 230 Cole, T., 277 Coleman, C., 232 Collinge, J., 208–209 Collins, S. J., 210 Collins, S. R., 174 Colman, D. R., 213 Colombo, G., 230 Colombo, L., 207, 213 Colon, W., 278 Come, J. H., 210 Condron, M. C., 210 Condron, M. M., 233 Connolly, J. L., 118 Connors, L. H., 120 Conroy, M. J., 281 Conway, J. F., 91, 94, 118, 177 Conway, K. A., 230 Cook, K. D., 282 Cooke, R., 179 Cooper, G. J., 175, 230–231 Cooper, G. S., 231 Cooper, T. G., 128, 174 Cordell, S. C., 63, 70, 86, 92 Cordes, H., 206 Corey, R. B., 3, 14, 49, 98, 122, 156, 177, 211 Cornwell, G. G. III, 282 Coschigano, P. W., 136, 174 Cosgrove, L. J., 92 Cossart, P., 92 Costa, P. R., 14, 20, 278 Costas, C., 119 Costello, C. E., 120

Cotman, C., 273 Cotter, R., 278 Cotter, S. E., 15 Coulary-Salin, B., 172–173, 175, 177, 272, 274 Courvalin, P., 93 Cousens, S. N., 206, 213 Coustou, V., 127, 131, 174 Coustou-Linares, V., 138–139, 174 Cowen, L., 91 Cox, B. S., 167, 174, 177 Craig, C. L., 18, 23, 31, 35, 39, 44, 47, 50 Craven, C. J., 281 Crestfield, A. M., 274 Cross, T. A., 43 Crowell, R. L., 118 Cullin, C., 178–179, 277 Cunningham, J. A., 118 Curiel, D. T., 120 Currey, J. D., 44 Curtain, C., 210 Curti, B., 91, 96 Cusack, S., 118–119, 121, 123–124, 179 Cuzange, A., 120 Cygler, M., 92 Czerwinski, E. W., 62, 67, 92

D D’Ursi, A. M., 274 Da Costa, M., 213 Daggett, V., 207 Dagkesamanskaya, A. R., 179 Daidone, I., 207 Dal Degan, F., 94 Damas, A. M., 230 Dames, S., 213 Daniels, A. U., 233 Das, A. K., 211 Das, C., 206 Datki, Z. L., 277 Davidoff, M. R., 46 Davidson, W. S., 230 Davies, D. R., 173, 179, 206, 272, 277, 282 Davies, G. J., 91, 95, 122 Davies, M. C., 230 Davies, P. L., 94 Davis, E. A., 45 Dayan, S., 122 De Gioia, L., 213

288

AUTHOR INDEX

de Haard, H. J., 122 Dea, J., 51 DeArmond, S. J., 173, 176–177, 210, 212–213 Decatur, S. M., 279 Decottignies, P., 173, 273 Dedieu, J. C., 213 DeGioia, L., 207 Dehdashti, S. J., 62, 67, 92 Deisenhofer, J., 76, 93, 120, 123, 210 Delaglio, F., 14, 233, 279 DeLano, W. L., 57, 92, 101, 119, 274 Deleu, C., 174 Della Vedova, F., 213 DeMarco, M. L., 207 Demura, M., 43 Demuro, A., 230 Deng, F., 53 Deng, H. X., 274, 280 Denis, P., 282 Denny, M. W., 18, 20–21, 44 DePace, A. H., 152–153, 156, 164–165, 174 Derkatch, I. L., 128, 131, 166–167, 174 Dermody, T. S., 118, 121 Der-Sarkissian, A., 10, 13, 156, 159, 174 Desbruslais, M., 208 Desmyter, A., 122 Dessen, A., 123 Devaux, C., 99, 119 Deverin, S. P., 120, 206, 273 de Vries, R., 42 Di Nola, A., 207, 230 Diaz, F., 42 Diaz-Avalos, R., 136, 140–142, 146, 155, 161, 163–164, 168, 174–176, 207 Dickerson, R. E., 277 Dickinson, S., 272 Dickmanns, A., 95 Dicko, C., 17–18, 20, 26–31, 34–36, 44–45 Dickson, R. C., 114, 123 Dideberg, O., 94 Diehl, T., 231 Diekmann, S., 275 Dijkstra, B. W., 96 Dill, K. A., 35, 37, 39, 45 Diomede, L., 207, 213 Diprose, J. M., 118–119 Dische, F. E., 274 Divry, P., 230, 274 Do, K. G., 46

Doan, C. N., 92 Dobeli, H., 232 Dobson, C. M., 27, 35–37, 39, 41, 44–45, 120–121, 144, 174, 176, 179, 211, 230–232, 273–277 Dodatko, T., 58, 63, 68, 70, 82, 86–87, 92 Dodson, E. J., 95, 122, 208, 272 Dodson, G. G., 208, 272 Doey, L. J., 208 Doi, Y., 52 Domenici, P., 52 Donald, A. M., 27, 31, 45, 232 Donaldson, D., 280 Dong, A., 13, 174, 274 Donlan, M. E., 122 Donne, D. G., 207, 209 Dos Reis, S., 144–145, 150, 172–173, 175, 177, 272, 274 Doucet, D., 94 Doucet, J., 173, 273 Doucette, P. A., 275 Doudevski, I., 233 Dover, S. D., 206 Downie, A. R., 13 Downing, A. K., 273 Downing, D. T., 28, 48, 84, 94, 116, 120 Doyle, J. P., 213 Drew, M. G. B., 211 Driscoll, P. C., 273 Droguett, G., 118 Drouet, E., 119 Drueckhammer, D. G., 91 Drummond, D., 206 Dubochet, J., 147, 175 Dubois, S., 173, 175, 273, 275 Duffy, L. K., 207, 210, 275, 277 Dunaway, D. L., 28, 52 Dunham, C. M., 206 Dunker, A. K., 146, 175, 274 Dunwell, J. M., 73, 92 Durand, F., 178 Durmort, C., 99, 119 Durrenberger, M., 233 Duvezin-Caubet, S., 177 Dwek, R. A., 212 Dyda, F., 13, 272, 279 Dyson, H. J., 207 Dzwolak, W., 231

AUTHOR INDEX

E Eanes, E. D., 3, 13, 208, 230, 274 Eby, R. K., 18, 45, 49 Eck, M. J., 102, 119 Edmonds, B., 232 Edskes, H. K., 135, 138, 175, 178–180, 213 Edwards, C., 232 Efimov, A. V., 79, 92 Ehlerding, G., 275 Eichhorn, S. J., 53 Eigen, M., 207 Eisenberg, D. S., 12, 14, 92, 94, 206, 121, 144, 174, 177, 207, 209, 272, 276, 278, 280 Ekiel, I., 275 Eklund, H., 93 el-Agnaf, O. M., 206, 273 El-Ahmad, M., 93 El Khoury, J., 207 Elam, J. S., 275 Elleman, T. C., 92 Elliott, A., 13, 206 Emsley, P., 62, 67–68, 70, 85, 88, 92, 98, 113, 119 Emtage, S., 91 Endo, N., 233 Engel, A., 175, 230 Engel, J., 113, 119 Engh, R., 273 Engler, J. A., 113, 120 Erickson, B. W., 120, 122 Ernst, D., 13, 174, 274 Ernst, M., 14, 95, 178, 280 Eroshkina, A., 91 Esnouf, R. M., 180 Esposito, L., 94 Estermyer, J. D., 211 Esteve-Moya, V., 211 Estibeiro, K., 213 Evdokimov, A. G., 61, 92

F Fa¨ndrich, M., 275 Fahnestock, S. R., 31, 45 Fairbrother, W. J., 52 Fairweather, N. F., 92, 119 Fanchon, E., 94 Fandrich, M., 35, 37, 41, 45

289

Farina, L., 207 Farmer, B., 49, 51–52 Farr-Jones, S., 209–210 Fasman, G. D., 74, 91 Fassy, F., 95 Fay, N., 143, 146, 153, 175, 275 Fedic, R., 18, 25, 39, 45 Fedorov, A. A., 92 Felder, C. E., 178 Fender, P., 121 Feng, Z., 91 Fernandez, R. C., 95 Fernandez-Bellot, E., 179 Fernandez-Tornero, C., 9, 13 Ferrari, D., 96 Ferrero, C., 50 Ferrone, F., 207 Ferry, J. G., 93 Fezoui, Y., 233 Ficner, R., 95 Fields, B. N., 121 Figlewicz, D. A., 280 Finberg, R. W., 118, 122 Finch, J. T., 122, 178, 212, 279, 282 Fink, A. L., 45, 51, 146, 175, 335 Fiorito, F., 208, 211 Fischer, M., 173 Fischle, W., 213 Fisher, S., 282 Flaherty, K. M., 91 Flanagan, J. M., 118 Fletcher, M. A., 45, 275 Fletterick, R. J., 175, 177, 207, 209–211, 275, 279 Flock, D., 230 Floege, J., 275 Florencio, F. J., 96 Florkin, M., 230, 274 Fociani, P., 208 Foelix, R. F., 35, 45 Fontano, E., 174, 207 Foo, C. W. P., 21, 36, 45 Forge, V., 121–122, 172, 175, 211, 272, 274–275 Forloni, G., 207, 212–213 Formigli, L., 230 Forrest, J. C., 118 Forsyth, V. T., 50, 211, 121–122 Fossey, S. A., 31, 45, 49 Foster, D., 212

290

AUTHOR INDEX

Fournier, M. J., 48 Fox, G. C., 119 Frangione, B., 180, 233, 274–275, 280, 282 Frank, R. W., 213 Frank, S., 114, 119 Fraser, H., 206 Fraser, P. E., 207, 209, 213, 231, 233, 275, 276, 281 Fraser, R. D. B., 3–4, 13–14, 82, 92, 98, 119, 208 Fraser, R., 31, 45 Freddi, G., 49, 51 Freed, R., 232 Freeman, R., 175 Freimuth, P., 118 Frenkel, M. J., 92 Frey, P., 175, 230 Fridovich, I., 275 Friemann, R., 93 Frische, S., 38, 45, 50, 52 Frishman, D., 208 Fu, J., 213 Fuller, S. D., 118 Furst, F., 14, 95

G Gabizon, R., 207 Gabriel, J. M., 209 Gagnon, J., 121 Gajdusek, D. C., 178, 208, 280 Gal, L., 95 Galan, B., 14 Gallagher, K. S., 52 Galperin, M. Y., 96 Garb, J. E., 20, 38, 45 Garcia, E., 13 Garcia, J. L., 14 Garcia, P., 14 Garrett, T. P., 60, 92 Gaspert, A., 178 Gasset, M., 150, 175, 177, 211, 275, 279 Gast, K., 213 Gasteiger, E., 75, 92 Gattiker, A., 92 Gaub, H. E., 51 Gazit, E., 118–119, 122, 124, 212–213 Gebert, R., 176 Geddes, A. J., 18, 40, 45, 98, 119

Geiger, J. H., 93 Geisow, M. J., 273 George, J. M., 230 Gerard, R. D., 120, 123 Gerardy-Schahn, R., 95 Gerfen, G. J., 206 Gernert, K. M., 122 Getzoff, E. D., 274 Ghibaudi, E., 207 Ghidoni, J. J., 276 Ghiso, J., 233, 274 Giaccone, G., 208 Giampaolo, L., 213 Giannini, S., 281 Giannoni, E., 230 Giannoni, G., 211 Gibbs, C. J. Jr., 178, 280 Gibbs, N., 209 Gibson, B. W., 213 Gibson, K. D., 45 Gibson, T. J., 176 Gido, S. P., 50 Gielen, G., 123 Giese, A., 281 Gilbey, S. G., 274 Gilead, S., 118–119 Gill, A. C., 208, 211 Gilliland, G. L., 91, 96 Gimenez-Gallego, G., 13 Gingery, M., 209, 276, 280 Giorgetti, S., 278 Girola, L., 207, 213 Glabe, C. G., 179, 230, 232, 273, 281 Glass, N. L., 275 Glenner, G. G., 3, 13, 208, 212, 230, 274–275, 280 Gliozzi, A., 233 Glockshuber, R., 175, 212 Glover, J. R., 139, 141, 144, 147–148, 150–151, 155, 175, 177 Go¨rbitz, C. H., 208 Godzik, A., 92 Goeltz, P., 121 Goffredo, D., 208 Goldberg, E., 92, 121–122 Goldblum, R. M., 92 Goldie, K. N., 121, 175, 211, 230 Goldman, A., 94, 121 Goldsbury, C. S., 163, 175, 230–231, 233 Goodin, D. B., 213

291

AUTHOR INDEX

Gophna, U., 119 Gordon, D. J., 272, 275 Gorevic, P. D., 274–275 Gorla, S., 213 Gosal, W. S., 275 Gosline, J. M., 18, 20–21, 46 Gossert, A. D., 208 Goto, J., 280 Goto, Y., 231, 276, 278 Gotoh, Y., 51 Gotte, G., 275, 278 Gout, I., 273 Goux, W. J., 208–209 Govaerts, C., 10, 14, 116, 119, 129, 134, 150, 175, 208, 276 Gowda, D. C., 51 Gowland, I., 208 Gracz, H. S., 46 Grama, L., 277 Granja, J. R., 206 Grant, M. A., 210 Gray, V. T., 175 Grazia Bruzzone, M., 207 Greco, A., 14 Green, J. D., 230–231 Green, N. M., 102, 119 Gregory, D. M., 13, 173, 230, 273–274, 276 Greiner, R. A., 206 Gribskov, M., 76, 92 Griffin, R. G., 14 Griffith, J. D., 120 Griffiths, J. M., 10, 14, 120, 278 Grimes, J. M., 118–119 Griniuviene, B., 121 Gronenborn, A. M., 177 Gross, H., 176 Gross, M., 231 Grosse, F., 50 Groth, D. F., 177, 207, 209–213, 279–280 Grothe, R., 14, 121, 174, 177, 206–207, 272, 278 Grover, R. K., 232 Groves, M. R., 59, 61, 92 Grubb, A., 276, 279 Grubb, D. T., 18, 21, 46 Gruijters, W. T., 175, 230 Gryczynski, Z., 215 Grynberg, M., 92 Grzeschik, K. H., 277 Grzonka, Z., 276, 279

Guardado Calvo, P., 98, 100, 103, 106, 119 Gueńebaut, V., 213, 282 Gueft, B., 276 Guerette, P. A., 46 Guerrini, R., 274 Guijarro, J. I., 120, 231, 276–277 Guilligay, D., 118 Guinier, A., 208 Guiroy, D. C., 208 Gulich, S., 15, 180 Gulik-Krzywicki, T., 213 Gunasekaran, K., 213, 273 Guntert, P., 206, 211 Guo, J. T., 10, 14, 276, 282 Guo, Y., 119 Guo, Z., 178, 233

H Ha, K., 213 Ha, S. W., 38, 46 Hagihara, Y., 276, 278 Hahne, S., 281 Hainfeld, J. F., 155, 180 Haire, L. F., 208 Haldar, D., 211 Halimi, M., 207 Hall, J. E., 232 Hallewell, R. A., 279 Halverson, K. J., 34, 46, 120, 276, 278 Hamalainen, E. R., 121 Han, Z., 51 Hanan, E., 210 Hanby, W. E., 13 Haner, M., 229 Hannah, J. H., 94 Hansma, H. G., 49 Harding, M. M., 272 Hardy, J., 231 Harini, V. V., 206 Harker, D., 277 Harper, J. D., 39, 46, 230–231 Harrah, T., 122 Harris, C. M., 51 Harris, R. D., 51 Hart, P. J., 275 Hart, R. G., 277 Hartley, D. M., 231, 233 Hartwig, M., 278

292

AUTHOR INDEX

Hasegawa, K., 93, 176, 231, 276–278 Hashemolhosseini, S., 114–115, 119 Hasnain, S. S., 275 Hauser, N., 231 Hayashi, C. Y., 18, 20, 38–39, 45–46 Hayes, S. F., 13, 174, 213, 274 Hayward, L. J., 275 Hecht, L., 211 Hecht, M. H., 36, 44, 112, 119, 122–123 Heddle, J. G., 95 Heegaard, P. M., 206 Heffron, S., 76, 92 Hegde, S. S., 58, 63, 67, 70, 86, 92 Hege, T., 63, 68, 70, 75, 85, 92 Heidelbach, F., 50 Heikenwalder, M., 178 Heim, G., 231 Heise, H., 231 Hendrickson, W. A., 213 Hendsch, Z. S., 120, 278 Hennetin, J., 58–61, 73–75, 77, 79–80, 82, 85, 92, 171, 175 Henning, U., 119 Henrissat, B., 94 Henry, L. J., 113, 120, 123 Henschen, A., 273 Hensley, P., 213 Hensman, J., 45 Hentati, A., 274, 280 Hepburn, H. R., 18, 40, 46 Herberstein, M. E., 47 Hermann, R., 229 Hermo Parrado, X. L., 119 Hermoso, J. A., 9, 14 Hernandez, J.-F., 122, 211 Herrmann, H., 151, 175, 231 Herrmann, T., 211 Herron, S. R., 95 Herskowitz, I., 178 Hertel, C., 231 Herzfeldt, B., 274 Heslot, H., 28, 31, 46 Hess, M., 105, 120 Hess, S., 43 Hetz, C., 174 Hewitt, E. W., 275 Heyes, C. D., 233 Hickman, S. E., 207 Higgins, L. D., 281 Higham, C. E., 231

Hijirida, D. H., 30, 46 Hilbich, C., 213, 276 Hill, A. F., 208 Hillner, P., 174 Hilschmann, N., 277 Hindennach, I., 119 Hirakura, Y., 231 Hiramatsu, H., 231 Hirao, J., 176 Hiraoki, T., 47 Hirota-Nakaoka, N., 231 Ho, J. G., 9, 14 Hodak, H., 91 Hodgkin, D. C., 272 Hoenger, A., 121, 211 Hofman, A., 213 Hofmann, K., 91 Hofnung, M., 123 Holtet, T., 52 Holzemann, G., 233 Homans, S. W., 278 Homo, J.-C., 175 Hong, J. S., 113, 118, 120 Hong, J. Y., 174 Hood, L. E., 281 Hood, L., 213 Hoogland, C., 92 Hope, J., 206, 208 Horan, R. L., 42 Horiuchi, M., 207–208 Hornemann, S., 134, 175, 208, 212 Horstedt, P., 279 Horwich, A. L., 208 Horwitz, M. S., 118 Hosemann, R., 208 Hoshino, M., 276 Hosia, W., 213 Hossain, K. S., 27–28, 46, 49 Hosszu, L. L., 209 Hotchkiss, A. T., 95 Houghten, R. A., 178, 212, 279 Hounslow, A. M., 281 Howley, P. M., 121, 180 Hoyer, W., 231 Hronska, M., 28, 30, 46 Hsu, M., 44 Hu, P., 274 Hu, X. W., 50, 52 Huang, G., 95 Huang, W., 62, 85, 92

293

AUTHOR INDEX

Huang, X., 210 Huang, Z., 177, 207, 209, 211, 279 Hubbard, R. E., 91 Hubbell, W. L., 280 Huber, A. E., 52 Huber, R., 14, 95, 122–123, 179, 273 Hudson, S. M., 46 Huff, M. E., 231 Hugel, T., 51 Hughson, A. G., 213 Hugouvieux-Cotte-Pattat, N., 93 Huiskonen, J. T., 121 Hung, W. Y., 274 Hunter, B. K., 44 Hunter, C. A., 209 Hunziker, P. R., 233 Hurle, M. R., 282 Hurshman, A. R., 276 Hutchinson, G., 103, 108, 120 Huxley, H. E., 151, 176 Hyde, C. C., 121 Hyman, P., 90, 92, 122

I Iakoucheva, L. M., 274 Iizuka, E., 20–22, 31, 34, 46–47 Iizuka, R., 210 Ikeda, S., 180, 282 Immel-Torterotot, F., 178 Inge-Vechtomov, S. G., 174 Inoue, S. I., 18, 20, 24, 40, 47, 51 Inoue, S., 47 Inoue, Y., 152–153, 175–176 Inouye, H., 112, 120, 173, 206–211, 213, 272–273, 275–279 Ionescu-Zanetti, C., 232 Ippel, H. J., 42, 279 Ippel, J. H., 276, 279 Iqbal, Z., 274 Ironside, J. W., 206, 213 Irving, T., 92 Isaacs, N. W., 92, 119 Ishida, H., 50 Ishida, T., 43 Ishii, Y., 13–14, 233, 272, 279 Isobe, I., 38, 47 Ito, S., 90 Itri, V., 213 Iussich, S., 207, 208

Ivanova, M. I., 209, 276 Ivanyi, I., 92 Iverson, T. M., 64, 67, 93

J Jackson, G. S., 209 Jackson, M., 276 Jacob-Dubuisson, F., 91, 93 Jacobson, D. J., 275 Jacrot, B., 119–120 Jaenicke, R., 35, 39, 47 Jaikaran, E. T., 231 Jakob-Rotne, R., 231 Jakuba, C., 42 James, T. L., 178, 207, 209–210, 212, 279 Janes, R. W., 31, 52 Janin, J., 173, 273 Jankowska, E., 276 Janowski, R., 276, 279 Jansen, R., 231 Jao, C. C., 13, 174 Jaquinod, M., 123 Jaroszewski, L., 92 Jarrett, J. T., 209 Jaskolski, M., 276, 279 Jass, J., 232 Jayakumar, R., 281 Jayasinghe, S. A., 10, 14, 156, 176 Jelinski, L. W., 18, 46, 53 Jenkins, J., 62, 67, 73, 77, 80, 83, 89, 93, 95, 159, 176, 276 Jerala, R., 276, 281–282 Ji, G. D., 21, 46 Jia, Z., 94 Jiang, Y., 153, 176 Jimeńez, J. L., 116, 120, 163, 176, 231, 277 Jin, H. J., 23, 25, 27, 35, 38, 47, 51 Jin, L.-W., 47 Jin, X., 64, 87, 93 Jobling, M. F., 210 Johansson, B., 282 Johansson, J., 11, 49, 213, 278 Johansson, K., 93 Johnson, T., 212, 279 Joiner, S., 208 Jonas, A., 230 Jones, E. M., 210, 232 Jones, J. A., 276

294

AUTHOR INDEX

Jones, S., 277, 281 Jones, T. A., 94 Jornvall, H., 93 Jovin, T. M., 231 Juehne, T., 15 Jullian, B., 92, 175 Jurnak, F., 15, 57–59, 73–75, 77, 83, 92, 94–96, 108, 120, 124, 159, 180, 213

K Kad, N. M., 232, 277 Kadler, K. E., 274 Kagan, B. L., 210, 231–233 Kahn, A., 277 Kahn, S., 282 Kajava, A. V., 1, 7, 10–12, 14, 55, 57–59, 61, 64, 67, 71, 75–76, 82–83, 85, 88–89, 92–94, 116, 120, 125, 149, 153, 156–157, 159–160, 175–176, 210, 232, 277, 282 Kalverda, A. P., 278 Kameda, T., 38, 43, 47 Kaminsky, W., 38, 47 Kammerer, R. A., 119 Kanamaru, S., 59, 72–73, 85, 93, 98, 110, 120 Kaneko, K., 209, 212 Kang, J., 277 Kanyo, Z. F., 210 Kaplan, D. L., 42–45, 47, 49, 51–52 Kapurniotu, A., 213 Karplus, K., 91 Karplus, M., 37, 39, 45 Karsai, A., 277 Karshikov, A., 273 Kartha, G., 277 Kasai, N., 49, 51 Kascsak, R. J., 173 Katagiri, G., 50 Kataoka, K., 233 Katou, H., 276 Kauppinen, S., 95, 122 Kauzmann, W., 37, 39, 47 Kawakami, T., 278 Kayed, R., 179, 230, 232, 281 Keen, J. N., 15, 95–96, 118, 120, 124, 180, Kehoe, L. E., 64, 67, 79, 87, 93 Keith, H. D., 211 Keller, G., 213 Kellermayer, M. S., 277

Kelly, J. F., 211 Kelly, J. W., 28, 35, 39, 47, 49, 210, 231–232, 276, 278, Kelly, S. M., 208 Kemp, J. A., 231 Kenaga, L., 278 Kenchington, W., 18, 50 Kendrew, J. C., 277 Kenne, L., 94 Kenney, J. M., 17, 25, 30, 35, 37, 40, 44–45, 47 Kerkam, K., 22, 47, 52 Kessel, M., 93 Kester, H. C., 95–96 Kheterpal, I., 282 Khurana, R., 232 Khuri, S., 92 Kikuchi, Y., 111, 120 Kim, U. J., 36, 47 Kim, Y. S., 92 Kimura, S., 47 King, C. Y., 175–176, 207 King, J., 50, 91, 107, 111, 118, 120, 122 Kirschner, D. A., 3, 5, 10, 14, 48, 52, 120, 173, 206–211, 213, 272–273, 275–279 Kirschstein, S. O., 51 Kiselev, N. A., 277 Kishimoto, A., 60, 93, 146, 160–162, 176, 277 Kishore, A. I., 47 Kishore, R., 43, 99 Kisker, C., 67, 69, 87, 93 Kisters-Woike, B., 276 Kistler, J., 175, 230–231 Kita, N., 95 Kitagawa, T., 210, 231 Kittler, M., 275 Kitts, P., 121 Kivirikko, K. I., 121 Klafki, H. W., 277 Kloareg, B., 94 Klunk, W. E., 277 Knauer, M., 273 Knaus, K. J., 210, 277 Knebel, D., 233 Knight, D. P., 18, 22–25, 31, 36–38, 44–45, 47–48, 50–53 Knight, M. M., 48 Knipe, D. M., 121, 180 Knowles, P. F., 118 Kobayashi, M., 47, 51

295

AUTHOR INDEX

Kobayashi, T., 90 Kobe, B., 58–59, 61, 64, 67, 76, 82–83, 88, 93–94, 176, 210 Koch, K. M., 277 Kocisko, D. A., 120, 210, 278 Kocsis, E., 94 Koehler, R., 212 Komar, A. A., 179, 277 Komatsu, K., 43 Kondraskina, E., 92 Konieczny, L., 280 Kopplin, L., 208 Korokhov, N., 120 Korth, C., 175, 210 Kos, J., 273 Kosmoski, J., 273 Kostyuchenko, V. A., 93, 108, 110–111, 120 Kourie, J. I., 210, 213 Kovesdi, I., 122 Kovoor, J., 25, 36, 48–49 Kowal, A. S., 175, 178 Koza, M. A., 50 Kozak, M., 276 Kozhukh, G. V., 278 Kramer, L., 119 Kranich, J., 178 Krasnykh, V., 114, 117, 120 Kratzin, H. D., 277 Kraulis, P. J., 210 Krebs, M. R., 232 Krejchi, M. T., 37, 48 Kreplak, L., 231 Kretzschmar, H. A., 281 Krishna, V., 232 Krishnan, R., 156, 161, 176, 278 Krithivas, A., 118 Kroll, M., 120 Kroon-Zitko, L., 282 KrUl, M., 280 Ksiezak-Reding, H., 155, 176 Kuhn, U., 210 Kuhnel, K., 63, 68, 94 Kummerlen, J., 43 Kurt-Jones, E. A., 118 Kushnirov, V. V., 174, 177, 179 Kusumoto, Y., 233 Kuzuhara, A., 43 Kwon, M., 273 Kwong, P. D., 124

L Laarmann, S., 15 Lacroix, E., 121, 211 Laemmli, U. K., 111, 120 Lai, Z., 232, 278 Lal, R., 232–233 Lambert, M. P., 232 Landwehr, R., 119 Lang, Y., 173 Langen, R., 10, 13–14, 84, 94, 156, 159, 174, 176–177, 179 Langs, D. A., 58, 94 Lankinen, H., 94 Lansbury, P. T. Jr., 46, 116, 120, 209, 231–232, 276, 278, 280 Lansbury, P. T., 14, 39, 46, 210, 230–231 Lantos, P., 208 Lario, P., 206 Larsen, S., 94–95, 122 Larsson, A. M., 62, 67, 94 Lascu, I., 172, 175, 272, 274 Lashuel, H. A., 232, 278 Latawiec, D., 177 Laue, T., 123 Laursen, H., 206 LaVerde, G., 53 Lavigne, G., 123, 179 Lawson, J. D., 274 Lazo, N. D., 28, 48, 84, 94, 116, 120, 210 Le Marechal, P., 173, 273 Leahy, D. J., 60, 91 Leak, K., 208 Leapman, R. D., 13–14, 172, 178, 229, 233, 272, 279 Leclerc, E., 212 Lee, B. I., 64, 67, 94 Lee, J. G., 122 Lee, J., 124 Lee, K. H., 23, 48 Lefevre, T., 50 Legname, G., 173, 176–177, 210, 212, 215 Leiman, P. G., 93, 120 Leinala, E. K., 64, 67, 70, 86, 88, 94 Lemaire, H. G., 277 Lemstra, P. J., 31, 48 Leo, J. C., 94 Lepault, J., 175 Lesk, A., 79, 122, 178, 212 LeVine, H. D., 218, 232

296

AUTHOR INDEX

Levine, M., 280 Levy, Y., 210 Levy-Nissenbaum, O., 119 Lewis, R. V., 18, 20, 38, 45–46, 51 Li Blatter, X., 231 Li, C. M., 47 Li, G., 40, 48 Li, H., 176 Li, L., 137, 176 Li, S. F. Y., 18, 48 Li, Y., 92, 122 Liberski, P. P., 208 Libonati, M., 275, 278 Lieber, C. M., 231 Liebman, S. W., 174 Lietzke, S. E., 60, 62, 70, 85–86, 94, 113, 120 Liivak, O., 53 Lillie, M., 46 Lilliehook, C., 281 Lim, A., 112, 120 Limido, L., 207 Lin, H., 233 Lin, M. C., 210, 232 Lind, M., 210, 277 Linding, R., 147, 176 Lindquist, S. L., 127–128, 131, 136, 138–139, 144–145, 156, 161, 176, 175–178, 278 Lindsay, A. M., 95, 122 Lindstrom, V., 279 Linhardt, R. J., 92 Liosatos, M., 232 Liou, Y. C., 58, 63, 67, 69–70, 75–76, 86, 88, 94 Lipman, D. J., 211 Lipp, H. P., 173 Liu, A., 180, 214, 282 Liu, H., 209–210 Liu, J. J., 175, 177 Liu, R., 278 Liu, Y. C., 26, 36, 48, 118, 278, 280 Lizonova, A., 122 Llamas-Saiz, A. L., 119 Llinas, M., 210 Lo Leggio, L., 64, 67, 69, 94 Lober, G., 51 Locht, C., 91, 93 Loeloff, R., 282 Lohner, K., 282 Loike, J. D., 207 Loll, P. J., 273

Lomakin, A., 40, 48, 233 Lombardi, S., 47 Long, C., 174, 207 Lopez de la Paz, M., 112, 121, 211 Lopez Garcia, F., 150, 177, 180, 211, 214, 282 Lopez, R., 13 Lorenzo, A., 232 Lorrain, D. S.-M., 208 Lorrain, P., 208 Lortat-Jacob, H., 100, 121 Lotz, B., 30–31, 48–49, 51 Lou, M., 92 Louis, J. M., 163, 177 Louis, N., 113, 121, 123 Lovrecz, G. O., 92 Lowe, J., 92 Lozez, A., 26, 48 Lu, H., 42 Lu¨hrs, T., 178, 180, 280 Luan, C.-H., 51 Lucas, F., 18, 25, 49 Luhrs, T., 14, 95, 211, 214, 232, 282 Luirink, J., 95 Lundgren, E., 276, 279, 281 Lundmark, K., 40, 49 Luo, Y., 282 Lupas, A., 82, 94 Lustig, A., 119 Luther, P. K., 281 Lutsch, G., 213 Lynn, D. G., 13, 156, 173, 177, 230, 273–274, 276 Lysek, D. A., 206, 208, 211 Lyubchenko, Y., 278

M Macdonald, T. R. R., 13 MacPhee, C. E., 117, 121 MacRae, T. P., 3–4, 13–14, 31, 45, 82, 92, 119, 208 Maddelein, M. L., 14, 95, 141, 145, 174, 177–178, 213, 280 Madsen, A. O., 14, 121, 177, 278 Madsen, B., 50 Maezawa, I., 47 Magasanik, B., 136, 174 Magoshi, J., 46–47, 49, 51 Magoshi, Y., 23, 26, 36–37, 39, 47, 49, 51

AUTHOR INDEX

Mahony, E. M., 124 Maillet, L., 178 Maiti, N. C., 232 Maji, S. K., 211 Makhov, A. M., 71–72, 83, 94, 120, 160, 177 Makin, O. S., 211, 278 Malesani, P., 212–213 Malinchik, S. B., 211, 278 Malisauskas, M., 232 Malone, M., 210, 277 Man, O., 178 Manabe, T., 43 Mancias, J. D., 123 Mangione, P., 278 Manning, J., 277 Mantsch, H. H., 276 Marcotte, E. M., 88, 94 Margittai, M., 84, 94, 159, 177 Markovic, O., 93 Marqusee, S., 210 Marrari, M. A., 213 Marrink, J., 274 Marsh, R. E., 18, 49, 211 Martıńez-Costas, J., 119 Martin, D. C., 42 Martin, J. D., 282 Martin, S. R., 118–119 Martinez-Ripoll, M., 14 Marusich, E. I., 121 Masison, D. C., 136, 145, 177, 278 Mason, P. E., 211 Mason, R. P., 211 Mason, T. L., 48 Massignan, T., 213 Masters, C. L., 180, 210, 276–278, 282 Mastrianni, J. A., 211 Masuch, R., 213 Matsui, T., 114, 121 Matsunaga, Y., 178, 212, 279 Matte, A., 92 Mattevi, A., 91, 96 Mattson, M. P., 178, 233 Maunsbach, A. B., 45 Maxwell, A., 92 May, B. C., 210 Mayans, O., 93 Mayer, F., 277 Mazzoleni, G., 213 McAllister, C., 278 McCardle, L., 206

297

McClain, R. D., 122 McColl, I. H., 211 McConnell, I., 206 McDonald, B. L., 278 McGhie, A. J., 48 McHolland, L. E., 212 McIntire, T. M., 179, 232, 281 McKay, D. B., 91 McKern, N. M., 92 McKinley, M. P., 212, 278, 280 McLachlan, A. D., 92, 119 McLachlan, D. R., 275 McLean, L. R., 279 McLeish, T. C., 118 McNeil, P., 180 McParland, V. J., 278 McPhie, P., 278 Mechling, D., 119 Meek, K. M., 274 Mehlhorn, I. R., 177, 207, 209, 211–212, 279 Meier, B. H., 14, 43, 46, 95, 178, 280 Meijer, H. E. H., 48 Melki, R., 146, 173, 175, 179, 273, 275, 277 Mendiaz, E. A., 282 Menendez, M., 14 Mengaud, J., 92 Menke, M., 91 Mercier, G. T., 114, 117, 121 Merckel, M. C., 94, 98, 102–103, 106, 112, 117, 121 Meredith, S. C., 13, 156, 173, 177, 230, 272–276 Merkle, H. P., 229 Merkle, M. L., 213 Merlini, G., 180, 278, 282 Merlino, A., 60, 94 Merz, P. A., 173 Mesyanzhinov, V. V., 93–94, 120–121, 123, 177 Meyer, R. K., 278 Michal, C., 46 Michel, G., 62, 67, 94 Michelitsch, M. D., 213, 282 Michikawa, M., 47 Midoro-Horiuti, T., 92 Miele, G., 178 Mierlo, v.C. P. M., 42 Migheli, A., 213, 274 Mikheeva, G., 120 Miller, L. D., 45 Miller, M. W., 212

298

AUTHOR INDEX

Miller, S., 95–96, 108, 114, 118, 122–123, 179 Miller-Auer, H., 13, 173, 230, 273, 276 Millhauser, G. L., 206 Millonig, R., 232 Milton, S. C., 179, 230, 232, 281 Mina, E., 230 Mini, T., 230 Minoura, N., 31, 49 Mironov, A. Jr., 134, 177 Miroshnikov, K. A., 114–115, 121 Mirzabekov, T. A., 210, 232 Missmahl, H. P., 278 Misur, M. P., 175, 230 Mitchenall, L. A., 92 Mitraki, A., 97, 102, 111–112, 117–119, 121–123, 179, 211 Miyazawa, M., 50 Mizuno, S., 47 Mizutani, K., 95 Mizzen, C. A., 275 Modler, A. J., 213 Moe, G. R., 92 Moeri, N., 91 Moineau, S., 122 Molinari, A., 213 Mollica, L., 213 Montag, D., 119 Monterroso, B., 14 Monti, M., 278 Monti, P., 31, 34, 43, 49, 51 Monticelli, L., 212 Monzani, E., 207 Moody, P. C., 95 Moore, A. M. F., 44 Moore, S., 274 Morbin, M., 213 Morera, S., 173, 273 Morgan, D. M., 274 Morgan, T. E., 232 Mori, H., 233 Morillas, M., 210, 277 Morita, T., 233 Moriyama, H., 213 Morozova-Roche, L. A., 232 Morris, E. P., 281 Morser, J., 213 Morten, I. J., 275 Moslehi, J. J., 178 Moss, D. A., 213 Moult, J., 82, 94

Mu¨ller, S. A., 175, 230 Muhlenhoff, M., 95 Muller, M., 50, 229 Muller-Hill, B., 277 Multhaup, G., 277–278 Muramoto, T., 129, 134, 177 Murphy, F. A., 99, 107, 121 Murray, I. A., 93, 209 Murzin, A. G., 91 Musco, G., 213 Musil, D., 273 Muthukumar, M., 37, 49 Mutter, M., 112, 121 Myers, S. L., 232 Myllyharju, J., 121 Myszka, D. G., 44

N Naber, N., 179 Nagashima, M., 213 Nagura, M., 49 Nagy, A., 277 Naiki, H., 231, 276, 278 Nakamura, S., 49, 51 Nakano, E., 47 Nakazawa, Y., 43, 53 Namba, K., 93, 176, 277 Narayanan, T., 213 Narizhneva, N. V., 51 Naumann, D., 213 Nawroth, P., 213 Nazabal, A., 140, 142, 177 Neduva, V., 176 Nelson, R., 12, 14, 116, 121, 144, 162, 177, 278 Nemethy, G., 45 Nemoto, N., 46, 49 Nentwig, W., 48 Nesloney, C. L., 28, 49 Nettleton, E. J., 176, 273, 277 Ng, D., 233 Ng, K. K., 14 Nguyen, A. D., 208 Nguyen, H. O., 176, 210 Nguyen, J. T., 177, 207, 211, 275, 279 Nibert, M. L., 102, 121 Nicholson, L. K., 43 Nielsen, E. H., 232

299

AUTHOR INDEX

Nielsen, K., 211 Nienhaus, G. U., 233 Nilsson, M., 279 Nitta, K., 43 Nivon, L. G., 211 Nodel, E., 95 Noppe, W., 232 Nordstedt, C., 281 Norstrom, E. M., 211 Novelli, A., 113, 121 Nowak, M., 280 Nowak, R. J., 232 Nummelin, H., 64–65, 67, 76, 85, 88, 94 Nusrat, A., 118 Nussinov, R., 213, 273 Nybo, M., 232

O O’Malley, M. B., 207, 275 O’Regan, J. P., 280 O’Rourke, K. I., 212 Oatley, S. J., 273 Ober, C. K., 49 Obmolova, G., 96 Obradovic, Z., 175, 274 Ochi, A., 28, 36, 46, 49 Oesch, B., 175 Ogawa, K., 43 Ohgo, K., 43 Ohkawa, Y., 47 Ohman, A., 279 Ohta, Y., 36, 43 Ohtomo, K., 47 Ohyama, E., 49 Okumura, T., 43 Olafsson, I., 279 Oliver, D. C., 70, 95 Olivieri, V., 175, 230 Olmstead, M. M., 206 Olofsson, A., 276, 279 Olsen, A., 49 Onnerfjord, P., 279 Oosawa, F., 211 Ooyama, E., 46 Orlova, E., 120, 231, 277 Orlowska-Matuszewska, G., 131, 177 Oroudjev, E., 23, 40, 49 Ortlepp, C. S., 46

Osanai, M., 43 Osherovich, L. Z., 136, 177 Osterroth, F., 14, 95 Otto, B. R., 63, 67, 70–71, 85, 95 Oudega, B., 95 Owen, D., 212, 279 Owen, R. J., 233 Oyler, N. A., 13, 174, 272, 274

P Padden, F. J. Jr., 211 Pakkanen, O., 114, 121 Pan, K. M., 157, 177, 207, 210–211, 279 Panine, P., 50 Papanikolopoulou, K., 97, 107, 113–114, 117, 121–122, 211 Pardowitz, I., 277 Parfenov, A. S., 173 Parge, H. E., 279 Park, J. Y., 47 Park, S. Y., 95 Parker, I., 230 Parker, K. D., 45, 119 Parker, T. M., 51 Parkos, C. A., 118 Parry, D. A. D., 1, 14, 82, 91, 119 Paternostre, M., 213 Patino, M. M., 138, 175, 177 Patskovsky, Y., 92 Pattanaik, A., 51 Patterson, D., 280 Patterson, J., 123 Pauling, L., 3, 14, 49, 98, 122, 156, 177, 211 Paushkin, S. V., 139, 177 Payton, M. A., 91 Peakall, D., 36, 49 Pearson, W. R., 211 Pedrotti, B., 213 Peisach, J., 206, 230 Pellegrini, M., 94 Pelton, J. T., 279 Peneff, C., 95 Penke, B., 277 Pepys, M. B., 176, 233, 274, 279, 281 Perera, L., 120 Peressini, E., 213 Peretz, D., 129, 134, 178, 206, 212, 279 Perrett, S., 172, 176, 180

300

AUTHOR INDEX

Perugini, M., 210 Perutz, M. F., 116, 122, 160, 178, 212, 279 Peters, H. M., 36, 49 Peters, P. J., 177 Petersen, T. N., 62, 67, 85, 95, 98, 113, 122 Peterson, S. R., 274 Petkova, A. T., 10–11, 14, 164, 178, 233, 272, 279 Petoukhov, M. V., 96 Petre, B. M., 232 Petsko, G. A., 206 Pettegrew, J. W., 277 Petty, S. A., 279 Pezolet, M., 50 Phillips, D. C., 277 Pick, A. I., 277 Pickersgill, R. W., 62, 67, 73, 77, 80, 83–85, 93, 95, 159, 176, 276 Picone, D., 274 Piekarska, B., 280 Pierce, M. M., 147, 149, 151, 178, 180 Pierce, N. E., 44 Pinilla, C., 178, 212, 279 Pitkeathly, M., 118 Plakoutsi, G., 233 Pleasance, S., 95 Pocchiari, M., 213 Podlisny, M. B., 207 Polavarapu, P. L., 213 Poli, G., 207 Pollard, H. B., 229 Pope, B. J., 212, 279 Porat, Y., 212 Portelius, E., 282 Porter, D., 18, 30, 39, 49, 51–52 Poser, S., 213 Poulsen, P., 94 Powell, S., 47 Powers, E. T., 276 Preiss, J., 93 Prelli, F., 274, 280 Prevelige, P. E. Jr., 107, 122 Price, N. C., 208 Prilusky, J., 147, 178 Principe, S., 278 Priola, S. A., 210 Prockop, D. J., 113, 119 Prota, A. E., 118 Proudfoot, A. E., 91

Prusiner, S. B., 14, 37, 49, 119, 127, 173, 175–178, 180, 207–213, 215, 275–276, 278–282 Ptitsyn, O. B., 27, 49 Pucci, P., 278 Puchtler, H., 280 Puoti, G., 208 Purvis, A., 92 Putthanarat, S., 18, 40, 45, 49 Pye, V. E., 64, 87, 95

Q Qu, Y., 115, 122 Quinn, T. P., 122 Quist, A., 233

R Race, R. E., 207, 213 Rachel, R., 122 Radford, S. E., 118, 173, 232, 275, 277–278, 281 Raetz, C. R., 60, 95 Rafferty, J. B., 93 Ragg, E., 212–213 Rahbar, F., 278 Rajagopal, K., 118, 122 Ramirez-Alvarado, M., 15, 180 Ramponi, G., 44, 174, 230 Ravasio, S., 96 Ravikumar, K., 58, 96 Raymond, G. J., 210, 212–213 Raymond, L. D., 207, 212 Reches, M., 122, 124, 212–213 Redeker, V., 173, 175, 273, 275 Reed, J., 13, 172, 272, 276 Rees, D. C., 93 Regan, L., 15, 180 Reichlin, T., 233 Reid, M. C., 51 Reinemer, P., 95, 122, 179 Reixach, N., 213 Relini, A., 233 Rennie, J. A., 274 Rerat, B., 273 Rerat, C., 273 Rhee, S., 179, 282

301

AUTHOR INDEX

Rhie, A. G., 211 Richardson, D. C., 70, 95, 112, 122, 280 Richardson, J. S., 70, 95, 122, 280 Richmond, J., 42 Ridley, S. P., 156, 178 Riedel, D., 231 Riek, R., 14, 95, 172, 175, 177–178, 180, 211–212, 214, 232, 272, 280, 282 Riekel, C., 14, 18, 20, 40, 44, 50, 177, 211, 121–122, 278 Riek-Loher, D., 232 Riesner, D., 176, 210 Rigby, A. C., 210 Rimmer, B., 272 Ripaud, L., 139, 145–146, 178 Rishpon, J., 124 Ritchie, M. A., 211 Ritter, C., 10–12, 14, 85, 95, 146, 160, 162–163, 172, 178, 232, 272, 280 Rizzo, N. W., 13, 172, 272 Robert, B., 213 Roberts, B. T., 180, 213 Roberts, C. J., 230 Robinson, C. V., 176, 273, 277 Robson, R. L., 95 Robson, R. M., 38, 50 Rochet, J. C., 280 Rockah, L., 119 Rocker, C., 233 Roderick, S. L., 60, 91–92, 95 Rodriguez, J. A., 275 Rodziewicz-Motowidlo, S., 279 Roelvink, P. W., 100, 122 Roher, A., 213 Rojas, E., 229 Rolandi, R., 233 Roller, P. P., 178 Romero, A., 13 Ron, E. Z., 119 Roos, R. P., 274 Rosano, C., 233 Rosch, P., 213 Rosen, D. R., 280 Ross, E. D., 135, 156, 178, 180 Ross, P. D., 173 Ross, S., 282 Rossi, G., 208, 213 Rossle, M., 23, 50 Rossmann, M. G., 83, 93, 95, 120, 123 Roterman, I., 280

Rougvie, M. A., 273 Rousseau, M. E., 31, 50 Routzahn, K. M., 92 Roy, R., 232 Rozenshteyn, R., 178, 212, 279 Rozovsky, I., 232 Rozwarski, D. A., 92 Rubenstein, R., 173 Rudall, K. M., 18, 50 Rudd, P. M., 212 Rudolph, R., 213 Ruigrok, R. W. H., 103, 120, 122, 211 Rupnik, M., 14 Russell, R. B., 176 Russell, W. C., 119 Rutkofsky, M., 208 Rybarska, J., 280 Rydberg, E. H., 178

S Saa, P., 174 Saafi, E. L., 175, 230 Sabourin, M., 173 Saderholm, M. J., 120 Safar, J., 150, 178, 213, 280 Saibil, H. R., 120, 176, 231, 277, 280 Sailer, A., 206 Saito, H., 43 Sakaguchi, R., 43 Salbaum, J. M., 277 Salemme, F. R., 3, 14, 64, 95 Salmona, M., 207, 212–213 Salnikov, V. V., 173 Salvadori, S., 274 Salvo, H., 232 Sambashivan, S., 280 San Martin, C., 118 Sanders, G. H., 230 Sanders, J. K. M., 44, 209 Sanger, F., 280 Sangren, O., 281 Santoso, A., 174 Sapede, D., 21, 39, 50 Saper, M. A., 280 Sapp, P., 280 Saraiva, M. J., 180, 230, 280, 282 Sarma, R., 275 Sasson, S. B., 207

302

AUTHOR INDEX

Sauder, J. M., 91 Saupe, S. J., 14, 95, 131, 140, 172–175, 177–178, 272, 274, 280 Savage, K. N., 46 Savistchenko, J., 175, 275 Sawaya, M. R., 14, 121, 177, 209, 276, 278, 280 Sawicki, G. J., 178–179 Scavetta, R. D., 85, 94–95 Schafer, H., 43 Schatzl, H. M., 213 Scheibel, T., 153, 178 Scheraga, H. A., 45 Scherzinger, E., 212, 279 Scheuermann, T., 213 Schick, B., 96 Schiff, L. A., 121 Schindelin, H., 93 Schirmer, E. C., 175 Schleucher, J., 276 Schlichting, I., 94 Schlott, B., 50 Schlumpberger, M., 178 Schlunegger, M. P., 272, 280 Schmitter, J. M., 172–173, 177, 272 Schmitt-Ulms, G., 212 Schneider, J. P., 118, 122 Schnell, F. J., 118 Schoehn, G., 96, 119, 122–124, 211 Schorn, C., 208, 211 Schubert, D., 232 Schuler, B., 9, 14, 62, 67, 69, 95, 113, 122 Schulthess, T., 119 Schulz, B., 213 Schutz, M., 123 Schwarz, E., 213 Schwarz, P., 178 Schweins, R., 50 Scott, M., 177 Scott, W. G., 206 Scrocchi, L. A., 213 Seckler, R., 14, 35, 47, 95, 179, 122–123 Secundo, F., 44 Seeger, H., 135, 178 Seelig, A., 231 Seelig, J., 230–231, 233 Sehnal, F., 20, 23, 38, 45, 50, 53 Seidel, A., 53 Seifert, B., 178 Seitz, M., 51

Selkoe, D. J., 14, 207, 210, 231, 233, 277, 280 Selvaggini, C., 207 Separovic, F., 47 Serag, A. A., 280 Serban, A., 177, 211–213, 278–279, 282 Serban, H., 178, 212–213, 279 Serio, T. R., 146, 178 Serpell, L. C., 3, 13, 49, 121, 178, 211, 215, 233, 273, 278, 281 Seves, A., 44 Seydel, T., 50 Sezutsu, H., 20, 23, 50 Shamala, N., 206 Shanmugam, G., 213, 281 Shanmuganandam, V. D., 208 Shao, J. Z., 49 Shao, Z. Z., 38, 44, 48, 50, 52–53 Shapiro, L., 213 Sharma, D., 173, 206, 208–209, 213, 272 Shaw, J. T. B., 49 Shearman, M. S., 208 Sheat, S., 272 Sheiba, L., 51 Sherman, M. B., 277 Shevchik, V. E., 93 Shewmaker, F., 180 Shewry, P. R., 25, 51 Shinchuk, L. M., 213 Shindyalov, I. N., 91 Shipley, N. H., 46 Shirahama, T., 233, 281 Shiu, P. K., 275 Shivaprasad, S., 281 Shore, V. C., 277 Siddique, T., 280 Sidle, K. C., 208 Sieber, V., 92 Siemer, A. B., 14, 95, 178, 280 Sierks, M. R., 278 Sijbrandi, R., 95 Sikorski, P., 49, 211, 213, 278, 281, Silman, I., 178 Silveira, J. R., 213 Silverstein, S. C., 207 Sim, V. L., 213 Simms, G., 278 Simon, E. J., 120, 278 Simon, M. N., 91, 93–94, 118, 172–173, 175, 177, 207, 232, 272 Sinclair, A., 210, 277

AUTHOR INDEX

Singh, S., 232 Sipe, J. D., 180, 213, 233, 282 Sirichaisit, J., 18, 50, 53 Skarzynski, T., 91 Skehel, J. J., 123 Skurnik, M., 94 Slattery, T., 213 Sletten, K., 282 Smirnov, V. N., 177, 179 Smith, D. A., 232, 275 Smith, D. P., 232, 281 Smith, D., 59, 72–73, 82, 85, 93, 95 Smith, J. M., 281 Smith, N. L., 59, 72–73, 82, 85, 95, 122 Smith, P. G., 213 Smith, S. G., 49 Smits, M., 212 Snidwongse, J., 93 Snigireva, I., 44 Snijder, H. J., 96 Snow, A. D., 275 Soares, J., 49 Sodroski, J., 124 Sohn, S., 26, 39, 50 Sokolov, Y., 232 Sokolowski, F., 213 Solomon, B., 210 Somashekar, R., 51 Sommer, S. S., 178 Sondheimer, N., 128, 136, 139, 144, 179 Song, K. J., 208 Sonoyama, M., 31, 50 Soreghan, B., 273 Sorrentino, G., 274 Soto, C., 174 Spach, G., 49 Speed, M. A., 35, 50 Spencer, R. G., 120, 278 Speransky, V. V., 138–139, 173, 179 Spinelli, S., 110, 122 Sponner, A., 18, 23, 50 Sprang, S. R., 102, 119 Springer, K., 118 Squire, J., 151, 179 Sreerama, N., 26, 50 St Geme, J. W. III, 15 Stahl, N., 213 Stahlberg, J., 94 Staniforth, R. A., 281 Stefani, M., 44, 144, 174, 179, 230, 233

303

Stehle, T., 118, 121 Stehlin, C., 119 Stein, W. H., 274 Steinbacher, S., 14, 72, 76, 85, 95, 98, 107–108, 113, 122–123, 160–161, 179 Steinbuchel, A., 52 Steipe, B., 95, 122, 179 Stetefeld, J., 91 Steven, A. C., 7, 14, 55, 57, 71, 75–76, 82–83, 88–89, 91–94, 118, 120–121, 159–160, 173, 175–179, 206, 210, 232, 272, 277, 281–282 Stewart, L. R., 210 Stewart, P. L., 274 Sticht, H., 213 Stierhof, Y. D., 119 Stoffler, D., 233 Stolz, M., 231, 233 Stopa, B., 280 Stork, M., 281 Strandberg, B. E., 277 Strange, R., 275 Street, A., 3, 13 Strelkov, S. V., 91, 123 Strey, H. H., 50 Stribeck, N., 49 Stuart, D. I., 118–119 Stummeyer, K., 59, 72–73, 85, 95 Su, H., 92 Subramaniam, V., 231 Subramanya, G., 51 Sucholeiki, I., 46 Sugino, R., 43 Suh, S. W., 64, 67, 94 Sulzenbacher, G., 64, 67, 69, 87, 95 Sun, Y. Y., 48 Sunde, M., 37, 50, 115, 120, 123, 231, 233, 276–277, 281, Sunderji, S., 175, 230 Supattapone, S., 213, 282 Surewicz, W. K., 207, 210, 232, 275, 277 Sussman, J. L., 178 Suttie, A., 206 Sutton, G. C., 118–119 Suwa, Y., 233 Suzuki, A., 90 Suzuki, E., 14, 119 Suzuki, H., 43, 93, 176, 277 Suzuki, M., 212, 279 Svehag, S. E., 232

304

AUTHOR INDEX

Svergun, D. I., 96 Sweat, F., 280 Swietnicki, W., 210, 277 Szela, S., 52 Szumowski, K. E., 211, 278

T Tabeta, R., 43 Taddei, N., 44, 174, 230, Taddei, P., 34, 43, 49, 51 Tagliavini, F., 207–208, 212–213 Taguchi, H., 93, 175–176, 277 Tainer, J. A., 274, 279 Takiff, H. E., 92 Tame, J. R., 95 Tanaka, A., 281 Tanaka, K., 47 Tanaka, M., 141, 146, 167, 169, 179 Tanaka, N., 121 Tanaka, T., 38, 47, 51 Tang, S. L., 48 Tao, Y., 114, 123 Taraboulos, A., 207 Tateishi, J., 210 Tatham, A. S., 25, 51 Taubes, G., 35, 51 Tavan, P., 281 Taylor, A. B., 275 Taylor, E. J., 95, 122 Taylor, K. L., 139, 141, 144–145, 149–151, 157, 173, 179, 213, 272, 281–282 Taylor, L., 213 Teeter, M. M., 206, 273 Teichmann, S. A., 91 Teixeira, S., 121 Tendler, S. J., 230 Tenidis, K., 213 Tennent, G. A., 176, 281 Teplow, D. B., 48, 210, 213, 231, 233, 281–282 Teplyakov, A., 74, 96 Ter-Avanesyan, M. D., 128, 136–137, 177, 179 Terenius, L., 281 Terry, A. E., 28, 36, 51 Terry, M. J., 178 Terzi, E., 231, 233 Thakur, A. K., 281 Thaler, F., 213 Thiele, U., 273

Thiyagarajan, P., 274 Thogersen, H. C., 52 Thomas, C. A., 207 Thomas, E. L., 49 Thomas, G. J. Jr., 119 Thomassen, E., 98, 108–109, 111, 123 Thompson, J. B., 49 Thomson, N. H., 173, 232, 275 Thomson, R., 180 Thornton, J. M., 103, 108, 120 Thual, C., 145, 147, 149, 179 Thyberg, J., 213, 281 Tian, M., 20, 38, 51 Tillinghast, E. K., 25–26, 51–52 Tingey, A. P., 95 Tirrell, D. A., 48 Tittmann, P., 176 Tjernberg, A., 281 Tjernberg, L. O., 213, 281 To¨ro¨k, M., 281 Tocilj, A., 94 Toedt, J., 96 Tomiyama, T., 233 Tompa, P., 146–147, 179 Tonelli, A. E., 46 Torchia, M., 212–213 Torok, M., 156, 179 Torrassa, S., 233 Torres, M. L., 213 Townley, M. A., 25, 36, 51–52 Tran, L. P. P., 44 Trommer, B., 232 Trotter, I. F., 206 Trus, B. L., 94, 177 Tsai, C. J., 213 Tsai, H. H., 213 Tsuda, H., 47, 51 Tsugita, A., 121 Tsukada, M., 20, 31, 49, 51 Tsuprun, V. L., 277 Tuite, M. F., 127, 177, 179 Tuma, R., 121 Turing, A., 30, 51 Turk, E., 281 Turk, V., 273, 282 Turkenburg, J. P., 95, 122 Turnell, W. G., 282 Tweedy, N. B., 122 Tycko, R., 13–14, 91, 156, 172, 174, 178–179, 213, 229, 233, 272, 274–275, 279

305

AUTHOR INDEX

Tyshenko, M. G., 94 Tzaphlidou, M., 274

U Uemichi, T., 273 Ulrich, A. S., 47 Ultrich, A. S., 43 Ulyanov, N. B., 209–210 Umemura, K., 43 Umland, T. C., 137, 149, 179, 282 Unger, E., 50 Unterbeck, A., 277 Uptain, S. M., 128, 164, 168, 179 Urban, V. S., 50, 274 Urry, D. W., 30–31, 34, 51 Urs, R. G. K., 21, 51 Uversky, V. N., 27, 29, 35, 39, 51, 146, 179 Uyama, A., 43

V Vainshtein, B. K., 213 Valentine, J. S., 275 Valery, C., 213 Valluzzi, R., 25, 28, 35, 38, 47, 51–52, 92 Van Beek, J. D., 46 van den Heuvel, R. H., 58, 63, 68, 79, 87, 96 van Pouderoyen, G., 62, 67, 96 van Raaij, M. J., 59, 72–73, 85, 96–99, 102–103, 107–112, 117–121, 123, 160, 179 van Rijswijk, M. H., 274 Vanoni, M. A., 91, 96 Vasisht, N., 208 Vasquez, S., 231 Vassar, R., 282 Vassilev, P. M., 231 Vecchio, G., 44 Veltel, S., 94 Vergara, J., 212 Vergnaud, G., 88–89, 91 Verma, C., 208 Verrips, C. T., 122 Vetting, M. W., 92 Vijayan, M., 272 Viles, J. H., 207, 213 Villeret, V., 91 Vincent, J. F. V., 44 Viney, C., 23, 27, 39, 44, 47, 51–52

Viola, K. L., 232 Virchow, R., 233, 282 Visser, J., 95–96 Vitagliano, L., 94 Vitali, J., 62, 67, 92, 96 Voelter, W., 213 Vogel, K. W., 91 Volitakis, I., 210 Vollrath, F., 17–18, 20–21, 23–25, 30–32, 36–39, 41, 43–53 von Schroetter, C., 180, 211, 214, 282 Vrielink, A., 206

W Waddon, A. J., 48 Wade, W. W., 49, 51–52 Wadell, G., 118 Wahle, E., 210, 213 Wakatsuki, S., 91 Wakayama, I., 208 Waksman, G., 15 Waldner, M., 213 Walker, V. K., 94 Wall, J. S., 71, 86, 91, 118, 155, 172–173, 175–176, 180, 207, 232, 272 Wallace, B. A., 31, 52, 58, 96 Wals, P., 232 Walsh, D. M., 231, 233 Walter, S., 179 Waltho, J. P., 209, 281 Walz, T., 232 Wang, D. I. C., 50 Wang, F., 213 Wang, H., 48 Wang, J., 10, 15, 107, 112, 123, 180 Wang, L. Y., 159, 180 Wang, W., 123 Wang, X., 279 Wanker, E. E., 212, 279 Ward, C. W., 92 Ward, I. M., 45 Ward, S., 114, 123 Warwicker, J. O., 18, 30, 52, 213 Watanabe, Y., 43 Waterfield, M. D., 273 Watkins, P. J., 274 Waugh, D. S., 92 Wawrzycka, D., 131, 177

306

AUTHOR INDEX

Weatherford, D. W., 3, 14, 64, 95 Weber, M., 213 Webster, P., 44, 174 Weinman, N. A., 278 Weintraub, A., 95, 122–123 Weissenhorn, W., 115, 123 Weisshart, K., 50 Weissig, H., 91 Weissman, J. S., 152–153, 164, 172, 174, 177, 179, 208 Weissmann, C., 173, 180, 206 Wells, T. N., 91 Wernstedt, C., 274 West, M. W., 112, 123 Westbrook, J., 91 Westermark, G. T., 49, 274, 279 Westermark, P., 49, 144, 180, 274, 282 Wetzel, R., 14, 44, 116, 123, 213, 233, 276, 281–282 White, A. R., 210 White, J. T., 276 White, M. A., 92 Whitson, L. J., 276 Whyte, S. M., 208 Wickham, T. J., 122 Wicki, P., 175, 230 Wickner, R. B., 10, 14–15, 93, 120, 127–128, 135–136, 145, 173, 175–180, 206, 210, 213, 272, 277–278, 281–282 Wider, G., 175, 180, 212, 214, 282 Wijmenga, S. S., 276, 279 Wild, A., 233 Wiley, C. A., 208 Wiley, D. C., 123, 280 Wilke, M. E., 120 Wilkinson, M. J., 230 Will, R. G., 206, 213 Willbold, D., 213 Wille, H., 14, 119, 150, 175, 177–178, 180, 208, 213, 276, 282 Willery, E., 91, 93 Williams, A. D., 44, 282 Williams, E. S., 208, 212–213 Williams, R. J. P., 52 Williams, R. W., 179, 281 Williamson, P. T., 46 Williamson, R. A., 177–178, 210, 212, 279 Wilson, D., 31, 35, 52

Windle, A. H., 27, 31, 45 Wingfield, P. T., 94 Winkler, D. C., 172–173, 206, 272 Winkler, S., 52 Winter, R., 231 Winter, S., 51 Wirtz, S., 175, 230 Wise, M. J., 25, 47, 52 Wittinghofer, A., 94 Wolfson, H., 273 Wolman, M., 282 Wonacott, A., 91 Wong, C. W., 275 Wong, S. S., 46, 231 Wood, S. J., 282 Woody, R. W., 26, 50 Woolfson, D. N., 117, 121 Wootton, J. C., 25, 52 Worboys, K., 93, 95 Work, R. W., 21, 52 Wormald, M. R., 212 Worthington, C. R., 213 Wright, P. E., 207, 213 Wrigley, N. G., 119 Wu, L., 213 Wu, P., 179, 281 Wu, S., 91 Wu¨thrich, K., 175–177, 180, 206, 208, 211–212, 214, 282 Wyatt, R., 124

X Xia, D., 99, 120, 123 Xie, X., 48, 53 Xiong, L. W., 208 Xu, J., 51 Xu, L., 102, 123 Xu, Y., 14, 276, 282

Y Yamane, T., 43, 90 Yan, S. D., 213 Yan, Y., 120, 122 Yanagihara, R., 208 Yanagisawa, K., 47

307

AUTHOR INDEX

Yanai, A., 207 Yang, F., 20–21, 31, 34, 48 Yang, J. T., 46–47 Yang, M., 20, 31, 34, 43 Yang, S. L., 212 Yang, X., 114, 124 Yang, Y. H., 52 Yang, Z. R., 147, 180 Yang, Z. T., 53 Yankner, B. A., 232 Yao, H., 48 Yao, J. M., 38, 42–43, 53 Yao, J., 43 Yao, W. H., 20, 34, 53 Yao, Z., 45 Yates, J., 273 Yau, W. M., 13, 174, 178, 233, 274, 279 Ye, C. P., 231 Yeates, T. O., 94, 275, 280 Yee, V. C., 210 Yemini, M., 118, 124 Yeo, H. J., 9, 15 Yoder, M. D., 7, 15, 57–59, 67, 73–77, 83, 92, 94, 96, 98, 108, 113, 120, 124, 156, 159, 180, 213 Yoshida, M., 93, 176 Yoshimizy, H., 43 Yoshizawa, F., 43 Yoshizawa, K., 47 Young, R. J., 31, 50, 53 Young, S., 213 Yu, T., 48 Yukuhiro, K., 20, 23, 50

Z Zagorski, M. G., 232 Zahn, R., 150, 177, 180, 211, 214, 282 Zamotin, V., 232 Zax, D. B., 46, 53 Zeev-Ben-Mordehai, T., 178 Zeidler, M., 213 Zeller, N., 178 Zemanek, G., 280 Zerovnik, E., 276, 282 Zhang, C., 232 Zhang, G. F., 180, 282 Zhang, H., 209 Zhang, S., 117, 124 Zhang, X. J., 180 Zhang, Y. B., 39, 53, 118 Zhao, L., 213 Zhou, H. J., 53 Zhou, J. M., 172, 176, 180 Zhou, L., 39, 53 Zhou, P., 20, 24, 36–37, 48, 52–53, 174 Zhu, H., 213 Zhu, L., 145, 176, 180 Zirwer, D., 213 Zong, X. H., 53 Zsurger, N., 206 Zubieta, C., 105, 124 Zurabishvili, T. G., 94, 177 Zurdo, J., 120–121, 211, 230–231, 273, 277 Zurovec, M., 20, 23, 38, 45, 50, 53 Zylberberg, I., 36, 48

SUBJECT INDEX

A A8A, 187, 195 A . See Amyloid- fibrils A 1–40, physicochemical studies, 227 A 1–42, 222 A 22–35, 198 a þ type conformation, 191 Acetyl-coenzyme A, 87 Acinous glands, 38 Acinous silk proteins, 33–34 ADE marker, 167 Adenovirus(es), 99, 105. See also Adenovirus fibers Adenovirus fibers head domain, crystal structure of, 99–101 morphology, 99 shaft domain, crystal structure of, 98, 102–107 Adhesins of pathogenic Gram-negative bacteria, 88 viral, 85, 90 ADP-glucose pyrophosphorylase, 87 AFP. See Antifreeze proteins a-helical coiled-coils, 87, 102 a-helical domains, in PrPC and PrPSc, 187 a-helices, 9, 34, 151 parallel, 182 Ala117, interaction of His111 and, 196–197 Alanine, 20, 25, 38 Alanine-rich domain, of prion cytotoxicity, role of, in turn conformation, 197–198 3F4 epitope concealed in PrPSc at heterodimer interface, 195–196 H1 domains, 194–195 His111 and Ala117 interaction, 196–197 intermolecular hydrogen binding by A8A, 195

quarter-staggered -chain assembly, 191–194 Alkaline protease from Pseudomonas aeruginosa, 57–58 Alzheimer’s A 25–35 domain, 197–198 Alzheimer’s -amyloid analogues, 184 Amino acid(s) C-repeat of, 5 crystalline blocks in short side-chained, 18 residues, 58 residues 246-527 of, 108 residues 333-341 of, 110–111 role of filament formation and sequence in composition of, 156–157 sequence motifs of -arcs and recurring conformations, 76–80 sequences of, bioinformatic analysis of, bacterial virulence factors, 71 Amylin20-29 domain, 184 Amyloid(s), 3, 4 cross- pattern in fibrils of, 115 defining properties, 144–145 diseases involving, 199 physical appearance of filaments of, 145 and sheet packing, 5 structural models for, 11 structural studies of, 10–12 Amyloid- (A ) fibrils EM studies, 221 SFM studies, 224–225 structural model for, 262–264 Amyloid deposits classes in, 218 endocrine hormones in, 218 Amyloid fibril(s), 39, 115, 218–219, 236–239 amyloidogenic core domains of, 198–199 amyloidogenic regions and structures of, 84–85

309

310

SUBJECT INDEX

Amyloid fibril(s) (continued) and cross- structure, 40–41 cross- structure of, 237 intermediate stages in assembly pathways of, 223–226 model of, 39–40 nucleation, 39–40 polymorphism, 219–223 prediction of amyloidogenic regions and structures of, 84–85 SFM to study morphology of, 224–226 structural models of. See Amyloid-like fibrils, structural models of structure of, formed by human amylin peptides, 218 surface-confined assembly pathway of, 224 synchrotron X-ray fiber diffraction patterns of, 238 in vitro assembled, morphology of, 221 water-filled nanotubes model for, 60 X-ray fiber diffraction patterns, 218 Amyloid filaments of fungal prions, structural models for -helical models, 159–162 superpleated -structure model, 157–159 Amyloid-forming peptides and lipid membranes, 228 Amyloid-like fibrils, 236–238 structural models of, classes of. See Amyloidlike fibrils, structural models of Amyloid-like fibrils, structural models of, 238 and fibril properties. See Fibril properties, and structural models gain-of-interaction model, 243, 245–256 natively disordered proteins, models of, 257–264 refolding models, 239–244 Amyloidogenic core domains, of amyloid fibrils, 198–199 X-ray diffraction study, 198–199 Amyloidogenic polypeptides, 184 Amyloidogenic proteins, diseases involving, 199 Amyloid oligomers, cytotoxicity mechanism of, 226–227 Amyloidosis, 217 of prion domains, filament formation and prion conversion role, 143–151 Amyloid peptides effect of solvents, 223

soluble forms of, 223 solvents effect on conformation, 223 Antheraea pernyi, 25 Antibiotic gramicidin, 57–58 Antibody 3F4, 187 epitope recognized by, 195–196 Antifreeze proteins (AFP), 76, 86 from Choristoneura fumiferana, 67 Tenebrio molitor, 58, 67 Antifreeze solenoids, 70 a-silk, 18 Asparagine, 74, 79 Asparagine-ladders, 59–60 Atomic force microscopy (AFM), 186 Autotransporter protein Hia, crystal structure of, 9 Avian reovirus S1133, 101–102 triple -spiral repeats identified in, 103

B Bacterial glutamate synthase, 79 C-terminal domains of, 58, 68 Bacterial transferases, 69 trimer formation of, 86–87 Bacterial virulence factors, bioinformatic analysis of amino acid sequences of, 71 Bacteriophage PRD1, 99, 105 P5 protein, 103 structure of proteolytic C-terminal fragment of, 112 Bacteriophage T4, 108, 152 Bacteriophage tail proteins, 72 -arcades, 59, 78, 82 -arcs, 58–59, 77 -chain assembly, quarter-staggered, 191–194 -crystallite orientation of, 3 in polyglutamine, 203–205 Betabellins, 112 BETAWRAP, 76 -fibrous fold, 98 -hairpins, 9, 77, 78, 251 -helical model, 159–161. See also Water-filled nanotube model for amyloid fibrils disadvantage of, 161–162 and prion variants, 170–171

SUBJECT INDEX

-helices, 58, 83, 98 Bioinformatic sequence analysis, 71, 88, 90 -keratin, mammalian, 1, 3 -keratin, pleated sheet structures in, 3 Bombyx fibers, 21 Bombyx mori, 21–22 fibroin, 37 model assembly in, 24 silks, 25 silks, secondary structure of, 38 Bombyx mori silk -sheet structure, 98 Bordetella pertussis, 56–57, 67 FHA of, 83–84 -roll, 56–58 -serpentine model. See Superpleated -structure model -sheets, inverse temperature transition to, 34 -sheets structures, 3, 219, 226 crystalline, 198 and X-ray crystallographic studies, 9–10 -silk(s), 4, 18. See also Silk fibers pleated sheet structures in, 3 sol-gel transition, 25–30 -silk(s), model assembly in, and enrichment for -structures -sheet enrichment, 23–25 fiber construction, 22–23 -solenoid(s), 7–8, 57–58, 60–61, 71 amino acid compositions of, 74 amino acid sequences of, 74–76 arc conformations in, 80 -arches in, 81 -strand in, 77 capping and bulging, 70–71 classification of, 69–70 cross-sectional shapes, 67–68 crystal structures of, 61 differences between sequences of singlestranded and triple-stranded, 80–82 3D structure of, 74–75 elongated shape and rigidity of, 85–86 folds, paths of, 87–89 groups, 65–66 handedness, 61–63 histogram of amino acid composition of, 75 inferred tendency of, 88 multistranded, 71–74 nonredundant set of -solenoid proteins, 61 oligomerization of, 87

311

polypeptide chain in, 77 relationships between handedness and twist, 66–67 ribbon diagrams, 74 strand-turn-strand motifs in, 76–77 structural features of, 58–59 trimers, projections of, 69 triple-stranded, 59 twist, 63–66 -solenoid proteins, evolution of homologous, 89 paths of -solenoid folds, 87–89 -solenoid shaft, bulging of globular domains from, 71 -solenoid structures and amino acid composition, 74 functional implications of regular, 86 functional implications of shape and rigidity, 85–86 left-handed, 64, 66–67 as oligomerization motifs, 86–87 perspectives of sequence-based prediction of, 82–84 right-handed, 62–63, 66–67 as scaffolds for multidomain complexes, 87 -strands antiparallel, 194–195 and apolar and polar residues, 75 fibrous polypeptides, 116 in hydrogen-bonded -sheets, 2 interactions with apolar side chains and short peripheral, 8–9 linkers connecting, 110 of O-type coils, 68 parallel/antiparallel arrangements in, 156 side chains of, 110 swapping of, 86 upright orientation of, 100 -structural fibrous folds, 7–9 -structural fibrous topologies, crystal structures, 3 -structural folds -helical formulation, 12 superpleated -structure, 12 -structures in fibrous proteins, 1–4 characteristics of, 4–6 crystallographic studies and, 7–9 -turns, 19–20, 34 types II and VIa, 103

312

SUBJECT INDEX

C Calcitonin, EM studies, 220–221 CAP. See Cyclase-associated protein CAR protein. See Coxsackievirus and adenovirus receptor protein Cation– interaction, 195 C-domain, of PrP, 135, 137, 141, 182 and filament formation, 143 CD spectroscopy. See Circular dichroic spectroscopy Cell toxicity, of amyloid deposits, 219 Cellular form of prion(s). See PrPc Chimeric fiber-fibritin, 114 Choline-binding domains, in pneumococcal virulence factor LytA, 9 Choristoneura fumiferana, 67, 86 Chrysopa silks, cross -structure of, 98 Circular dichroic (CD) spectroscopy, 188, 268–269 CJD. See Creutzfeld-Jakob disease Clostridium difficile, carbohydrate-binding domain of toxin A from, 9 Cocoon silk, Bombyx mori, 21 Conformations, natively unfolded, of fungal prions, 146–148 Congo red, 144, 218 Coxsackievirus and adenovirus receptor (CAR) protein, 100 Creutzfeld-Jakob disease (CJD), in humans, 132, 185 Cross- diffraction pattern, of amyloid fibrils, 267–268 Cross- pattern, 115 Cross- silks, 18 Cross- spine, 116 structure, 12 Cross- spine model of fibril formation, 248–252 of HET-s fibrils, 260 Cross- structure, 4 in amyloid fibrils, 184 in amyloids, 144 in -helical models, 157 chrysopa silks, 98 in fungal prions, 146 of polyalanine, 199–201 Crystallization and silk strength, 37–39 Crystal structures, for adenovirus fiber, 112

Crystal structures, of viral fibers adenovirus, reovirus, and phage PRD1 fiber head domains, 99–102 adenovirus, reovirus, and phage PRD1 fiber shaft domains, 102–107 morphology, 99 tailed phage fibers, 107–111 C-terminal domains. See also C-domain of bacterial glutamate synthase, 58, 68 of glutamate synthase, 58 Cyclase-associated protein (CAP), 70 Cylindrical -helix model, 257–258 Cystatin C, 3D domain swapping in, 252–255 Cystatin C amyloid angiopathy, 253 Cysteine, 79 Cytotoxicity, of oligomers of amyloidforming peptides and proteins, mechanism of, 226–227

D Dalfopristin, 87 3D domain swapping models, of fibril formation, 252–255 with cross- spine models, 255–256 Direct-stacking models, 246–248 Diseases, involving amyloids, 199 Disordered proteins, intrinsically, 257 DNA gyrase effect of protein, 86 inhibitor MfpA from Mycobacterium tuberculosis, 67–68 MfpA inhibitor of, 70 DNA mimicry, 86 DNA recombinant technology, 31

E E. coli K1 bacteriophage K1F endosialidase, 72 Electron microscopy (EM), for amyloid fibril polymorphism, 219–223 Endolysin, 9 from pneumococcal bacteriophage Cp-1, 9 Erwinia chrysanthemi, 57 pectate lyase from, 98 Eukaryotic RP2 proteins, 61, 89

SUBJECT INDEX

F Familial disease, 185–186 3F4 antibody, 187 109–112 epitope recognized by, 195–196 Feather keratins, 98 -sheet structure of, 98 FHA. See Filamentous hemagglutinin Fibril assembly missing cross- structure, 40–41 spinning versus growth, 39–40 Fibril formation, cooperative kinetics, 270 Fibrillar amyloid structure, 112 Fibrillar morphology, of amyloid fibrils, 265–266 Fibrillogenesis, 223 Fibril properties and structural models, relation of, 264–265 Congo red binding and birefringence, 269 cooperative aggregation kinetics of formation, 270 cross- diffraction pattern, 267–268 extreme stability, 271 fibrillar morphology, 265–266 increase in structure, 268–269 protofilament substructure, 266–267 self-association, 269–270 variable morphology, 267 Fibrils of mammalian prion protein, 10 Fibril stability, 271 Fibritin’s trimerization, 114 Fibroins, 18, 25 phase diagram studies of, 26 Fibrous proteins, -structures in, 1–4 characteristics of, 4–6 crystallographic studies and, 7–9 Fibrous proteins, stability, folding, and assembly of biochemical stability, 111–112 general folding strategies employed by natural -stranded proteins, 112–113 natural -stranded folds and amyloid structures, 115–116 role of globular domains in folding of triple -stranded, 113–114 specific chaperones involved in folding of triple -stranded, 114–115 Fibrous proteins, structure and sequence of repeats present in, 104–105

313

Filament formation, in fungal prions in amyloidosis of prion domains, role of prion conversion and, 143–151 conformational changes and, 149, 151 N- and C-domains role in, 143 prion domains role in, 139–141 in vitro-assembled filaments, 139 in vivo-assembled filaments, 137–139 Filamentous hemagglutinin (FHA), of Bordetella pertussis, 56–57 Filament polymorphism. See Polymorphism, in fungal prion filaments Flagelliform silk proteins, 34 Fourier transform infrared (FTIR) spectroscopy, 188, 268–269 FTIR spectroscopy. See Fourier transform infrared spectroscopy Fungal prions, 171–172 -strands in, parallel arrangement of, 156 cross- structures, 145 defining properties of amyloids and, 144–145 filament formation in. See Filament formation, in fungal prions filament polymorphism in, 163–166 filaments, handedness of, 153 filaments, mass-per-unit-length of, 155 filaments, polarity of, 151–153 functional domains of, 136–137, 148 [Het-s], 131–132 infection mechanism, 133, 135 natively unfolded conformations, 146–148 nomenclature, 130 [PIN], 128, 131 prion domains of, 129, 135–136 prion domains role in filament formation of, 139–141, 143 protease resistance and physical stability, 145–146 [PSI], 128, 131 relationship of mammalian (neurotoxic) prions to, 132–134 secondary structure estimates of, 150–151 [URE3], 128, 131 variants of, 166–171 in vitro filament formation, 139 in vitro filament role in infection, 141–143 in vivo filament formation, 137–139

314

SUBJECT INDEX

Fungal prions, structural models for amyloid filaments of -helical models, 159–162 superpleated -structure model, 157–159

G Gain-of-interaction models, 243, 245 Type III or 3D domain swapping models, 252–255 Type II or cross- spine models, 248–252 Type I or direct-stacking models, 246–248 Type IV or 3D domain swapping with cross- spine models, 255–256 Globular proteins crystalline, 3 denaturated, 3 Glues, 18 Glutamate synthase, C-terminal domain of, 58 Glycine, 20, 25, 38, 74, 102–103 Glycosylphosphatidylinositol (GPI), 187 GNNQQNY, cross- spine model for, 248–252 Gp5, T4 cell-puncturing device, 72–73 Gp11, 111 Gp12, 115 T4 short tail fiber, 72 Gp34, 108

H Haemophilus influenzae, 9 Hamster PrP primary structure of, 184 residue number, physical-chemical parameters, 190 Handedness, prion filaments, 152–153 H 2m. See Human 2-microglobulin H-bonded -strands, 59 H-bonds -structural, 88 intercoil, 58, 60, 68, 70 network of, 59 peptide group, 79 H1 domain, of prion protein, 187 reverse turn conformation in, 194–195 Heterodimer interface 3F4 epitope at, 195–196

Heterodimer model, 185 HET-s fibrils, structural model for, 259–260 [Het-s] prion, 131–132 -helical model, 162 nomenclature, 130 prion domains of, 129, 135–136 HET-s protein, 131–132 1,1,1,3,3,3-Hexafluoro-2-isopropanol (HFIP), 223, 227 His111, 195 and Ala117 interaction, 196–197 Human amylin, 227 EM studies, 220–221 fibrils, 224–226 growth of, on mica, 224 peptides, structure of amyloid fibrils formed by, 218 SFM studies, 225–226 Human 2-microglobulin (h 2m), cross- spine model for, 248–252 Human prion diseases, 185–186 Hyaluronate lyase helix, 110 Hyaluronidase HylP1, 73 Hydrogen-bonded sheets, 5 Hydrogen bonds, intermolecular, 195 in triple-stranded -helix domain, 110 Hydrophobic proteins, 21

I Iatrogenic CJD (iCJD), 185 IGFR. See Insulin-like growth factor receptor Infectious disease, 185–186 Insect silks, 98 Insulin in amyloid deposits, 218 protofilaments, 239–240 Insulin-like growth factor receptor (IGFR), 60–61 Intrinsically disordered proteins, 257

J JAM-A. See Junction adhesion molecule A Junction adhesion molecule A (JAM-A), 102

K K1F endosialidase, 73

315

SUBJECT INDEX

L Lepidoptera, silk fiber formation in, 22–23 Leucine-rich repeat (LRR) proteins, 61 Lipid membranes, and amyloid-forming peptides, 228 Lipopolysaccharides (LPS), 111 LPS. See Lipopolysaccharides LRR proteins. See Leucine-rich repeat proteins

M Mammalian prion. See Prion protein Mammalian reovirus fiber, 103 Mass-per-length (MPL) measurements, of proteins, 221 M-domain of Sup35p, 136 MfpA inhibitor of DNA gyrase, 70 Micelle formation, 23 Miller index, structure factor, 192 Models, for prion amyloid filaments -helical models, 159–162 superpleated -structure model, 157–159 Models, structural, of amyloid and amyloidlike fibrils, 238 gain-of-interaction model, 243, 245–256 natively disordered proteins, models of, 257–264 refolding models, 239–244 and relation to fibril properties. See Fibril properties, and structural models Molecular dynamics study, of PrP106-126, 196 MPL. See Mass-per-length (MPL) measurements MPL, of prion filaments, 155 Mycobacterium tuberculosis, 67–68 R-type solenoid of MfpA protein from, 86 Myoglobin fibrils, 241 Myoviridae, 107–108

N Natively disordered proteins, models of amyloid fibrils, 262–264 cylindrical -helix model, 257–258 HET-s fibrils, 259–260 other models for polyglutamine, 258–259 Ure2p fibrils, 260–262

N-domain, of PrP,135, 182. See also Prion domains and filament formation, 143 of HET-s, 137 Nephila cocoon silk and in, 21 silks, concentration effect in, 27 silks, model assembly in, 24 Nephila edulis effect of protein-protein interactions on, 28 silk gland proteins from, 34 structural stability of ampullate silk protein in, 29 Nephila safety-line silk, 21 NMR study, for PrP peptides, 196 Nucleation-dependent model, for PrPSc, 185, 187

O Oculopharyngeal muscular dystrophy (OPMD), polyalanine extension in, 202 Oligomer(s), 86 of amyloid-forming peptides and proteins, cytotoxicity mechanism, 226–227 channel-type conductivity jumps of, 227 cytotoxicity in amyloid, 229 globular, 226 patch-clamping studies of, 227 small, 226 Olofsson model for TTR, 247 OPMD. See Oculopharyngeal muscular dystrophy Overt repeats, 76

P Parallel -helix, 57 Parallel -helix model, for PrP rods, 243 Parallel-stacked hairpin-like structure, for A fibrils, 263–264 Parallel superpleated -structure, for Ure2p fibrils, 261–262 PDB. See Protein data bank Pectate lyase C from Erwinia chrysanthemi, 57 Pectate lyase from Erwinia chysanthemi, 98

316

SUBJECT INDEX

Peptides. See also PrP peptides amyloid, soluble forms of, 223 polyglutamine, 10 seven-residue, crystal structure of, 162–163 Pertactin virulence, 67 [PIN], 128, 131 [PIN] prion domains of, 129, 135–136 nomenclature, 130 Pneumococcal bacteriophage Cp-1, endolysin from, 9 Pneumococcal virulence factor LytA, choline-binding domains in, 9 Podospora anserina, 259 prions of, 128 Podoviridae, 107 Polarity, of prion filaments, 151–153 Polar residues bulky, 80 ladders of, 57 positions occupied by, 59 regions enriched in, 75 uncharged, in amino acid composition, 135 Polar-zipper -sheet formation, 204 Polyacrylamide gels, 145 Polyalanine cross- structure in, 199–202 extension in oculopharyngeal muscular dystrophy, 202 Polyethylene glycol, 23 PolyGln15, slablike morphology of, 203 Polyglutamine -crystallite in, 203–205 fibrils, cylindrical -helix model of, 257–258 peptides, 10 reverse turn conformation role in, 205 spine, 255 Polyglutamine-RNase A fibrils, 3D domainswapping cross- spine model of, 256 Polymorphism in amyloid fibrils, 219–223 assembly, 219 of yeast prion fibrils, 12 Polymorphism, in fungal prion filaments, 163–166 and growth rate of filaments, 165 prion variants and, 167–168 Polypeptide chain, solenoidal windings of, 7 Polypeptides, amyloidogenic, 184

Polysaccharide-binding -solenoids, 85 Powder X-ray diffraction, 198 P.69 pertactin, 68, 88, 98 N-terminal coil of, 70 – interaction, aromatic, 195 – stacking, 39 PRD1 bacteriophage, 98 fiber, morphology, 99 fiber head domain, crystal structure of, 98–107 Prespun silk, 25 from Bombyx mori, 30 proteins, 21, 23 secondary structure population in, 28 solutions of, 31 stability and solubility of, 25 stored in, 26 transformation of, 23 Prion(s), 127. See also Fungal prions cellular form of. See PrPc conversion. See Prion conversion domain. See Prion domains scrapie form of. See PrPSc Prion alanine-rich domain cytotoxicity, role of, in turn conformation, 197–198 3F4 epitope concealed in PrPSc at heterodimer interface, 195–196 H1 domains, 194–195 His111 and Ala117 interaction, 196–197 intermolecular hydrogen binding by A8A, 195 quarter-staggered -chain assembly, 191–194 Prion and amyloid fibrils, structural models of gain-of-interaction models, 12 natively disordered models, 12 refolding models, 12 Prion conversion, 127, 132 in amyloidosis of prion domains, role of filament formation and, 143–151 Prion diseases classification of human, 185–186 Prion domains, 129, 135–136 amyloidosis of, 143–151 role in filament formation of fungal prions, 139–141, 143 Prion fibrils, 3–4 structural studies of, 10–12

317

SUBJECT INDEX

Prion filaments, structural models for, 11 Prion filament structure, constraints on dependence of filament formation on amino acid composition, 156 EM images of filament, 156 handedness, 153 mass-per-unit-length, 155 parallel arrangement of -strands, 156 polarity, 151–153 Prion pathology prion without genome, 186–187 PrPSc and PrPc, structural difference between, in 90–145 domain, 187–188 Prion protein (PrP), 182. See also PrPc; PrPSc fibrils of, 10 model for, 241, 243 nomenclature, 130 N-terminal fragments of, 10 prion domains of, 129 Prion-related peptides, 196 Prion variants, 166 detection methods, 167 and filament polymorphism, 167–168 superpleated -structure and -helical models and, 169–171 in vitro and in vivo filaments and, 169 Proline, 74 Protease resistance, of fungal prions, 145–146 Protein(s) intrinsically disordered, 257 simple -structural components of, 4 solenoid twist, 64 structurally reactive, 37 structures, 60 Proteinaceous infectious particle. See Prions Protein data bank (PDB), 61 Protein-protein interactions on Nephila edulis, effect of, 28 Proteins models, of natively disordered proteins amyloid fibrils, 262–264 cylindrical -helix model, 257–258 HET-s fibrils, 259–260 other models for polyglutamine, 258–259 Ure2p fibrils, 260–262 Protofibrils, 221 PrP. See Prion protein PrP106-126, 197

PrPc, 182 structural difference between PrPSc and, 187–188 PrPcwd, 183 PrPcwd-induced conversion, in cell-free system, 189 PrP27-30 domain, 188 PrP peptides, 10 NMR study for, 196 X-ray diffraction patterns of, 191–192, 194 PrP-related polyalanine, 10 PrPSc 130, 132, 182 3F4 epitope concealed in, 195–196 nucleation-dependent model for, 185, 187 relationship of fungal prions to, 132–134 structural difference between PrPc and, 187–188 PrP sequences pairwise sequence comparison of, 189–191 Pseudomonas aeruginosa, 57–58 [PSI] prion, 128 crystal structure of seven-residue peptide from, 162–163 genetic selection criteria for, 131 nomenclature, 130 prion domains of, 129, 135–136 variants and filament polymorphism, 167–168 Pyriform silk proteins, 33–34

Q Quarter-staggered -chain assembly, 191–194

R Refolding models, 239 for insulin protofilaments, 239–240 for myoglobin fibrils, 241 for PrP, 241, 243 for SH3 fibrils, 241–242 Reoviruses, 105 and bacteriophage PRD1 fiber C-terminal domains, 106 Reovirus fibers head domain, crystal structure of, 99–101 morphology, 99

318

SUBJECT INDEX

Reovirus fibers (continued) shaft domain, crystal structure of, 98, 102–107 Reverse turn conformation in H1 domain, 194–195 His111 and Ala117 interaction role, 196–197 role in polyglutamine, 205 Rhamnogalacturonase A, 98 Ribonuclease A (RNase A), 3D domain swapping with cross- spine models for, 255–256 RNase A. See Ribonuclease A Rnq1p protein, 128, 131 [RPSþ], 136 R-type -solenoids, 67–68 Run-away 3D domain-swapping, 253

S Saccharomyces cerevisiae, prions of, 128 Safety-line silks, Nephila, 21 Salmonella typhimurium, 9 Scanning force microscopy (SFM), for amyloid fibrils, 224–226 Self-association, in fibril formation, 269–270 Serine, 25, 79 Serralysin, 56–57 SFM. See Scanning force microscopy SHa90–231, disordered 90–145 domain within, 188 SH3 fibrils, refolding model for, 241–242 Silk-based peptide, 31 Silk fibers, 18, 23–25 amino acid, 39 composition of, 25 crystallization, 34 with different functions, 34 fibroin, formation of, 23 formation in lepidoptera, 22–23 formation in spiders, 22–23 formation of, 23–25 glands produce multiple proteins, 23, 25 inverse temperature, 34 mechanical properties, 18 and optical spectroscopy, 31 phase diagrams, 39 production process, 41–42

safety-line, 21 strength and crystallization, 37–39 structure and function relationship in, 18–21, 34–35 X-ray diffraction studies and, 18 Silk fibroins, 98 Silk protein(s), 18 acinous, 33–34 impact of SRCD on, 31 solubility of, 26 structure function relationship in, 19–20 structures, 30 study of problems in, 30 Silk protein(s), role and function of -sheet assembly in -transition, 31–35 factors governing transition to -structures, 35–37 irregular sequence units and heterogeneous structures, 37–39 monitoring and studying behavior, 30–31 Silk worms. See Bombyx mori; Nephila edulis Siphoviridae bacteriophage, 107 Siphoviridae proteins, 110 Slablike- structures, 184 SOD. See Superoxide dismutase Solenoidal windings of polypeptide chain, 7 Solenoid proteins, 58. See also -solenoid(s) Solvents, effect of, on amyloid peptides, 223 Spherulite, 38 Spider(s) silk, natural evolution of, 25 silk, fiber formation in, 22–23 silk gland distribution from orb weaver, 32 spinning silks, 31 Spidroins, 18, 25 time-induced conformational change of, 27 Spikes, 108 Spiral -hairpin stacks, 7–8 staircase, 9 Sporadic disease, 185–186 SRCD. See Synchrotron radiation-based circular dichroism Streptococcal phage-encoded hyaluronidase HylP1, 72 Streptococcal prophage SF370, 110

319

SUBJECT INDEX

Structural models for fungal prions. See Models, for prion amyloid filaments of fibrils. See Models, structural, of amyloid and amyloid-like fibrils Superoxide dismutase (SOD), directstacking model for, 246–248 Superpleated -structure, 12 Superpleated -structure model, 157–159 and prion variants, 157–159, 169–170 Sup35NM filaments, 135 Sup35p, 128. See also [PSI] prion -helical model for filaments of, 160–161 fibrils, 257–258 M-domain of, 136 protease resistance, 145–146 SurfD, 70 Synchrotron radiation-based circular dichroism (SRCD), 31 Synechocystis sp., 58 Synthetic polypeptides, 3 a-Synuclein, 10, 156, 159 pH in, 226 physicochemical studies, 227 Syrian hamster 90–231 (SHa90-231) prion, 188

T Tachykinin family, of neuropeptides, 198 Tailed bacteriophages, division of Myoviridae, 107 Podoviridae, 107 Siphoviridae, 107 T4 cell-puncturing device gp5, 72 Temperature-sensitive mutation, 115 Tenebrio molitor antifreeze protein from, 58, 67, 75, 86, 88 R-type -solenoid of antifreeze protein from, 69 T4 gp12, 73 ‘‘Thermodynamic hypothesis,’’ of Anfinsen, 245 Threonine, 74, 79 TNF. See Tumor necrosis factor Transmissible spongiform encephalopathies (TSE), 132 variants, 167 Transthyretin (TTR), EM studies, 220

Triangular cross -prisms, 7–8 Trimeric P22 tailspike, 98 Triple -spirals, 7–9, 102–103 in adenovirus fiber shaft, 107 Triple -spiral topology, 198 Triple -stranded folds mechanism evolved in formation of, 114–115 role of globular domains in formation of, 113–114 Triple-stranded -helix, 110 Triple-stranded -solenoids, 7–8 Trp, 70 TS -solenoids, 73 TSE. See Transmissible spongiform encephalopathies T4 short tail fiber, 72–73, 98, 109 TTR. See Transthyretin (TTR) fibrils TTR fibrils, direct-stacking model for, 246–248 T-type -solenoids, 67 Tumor necrosis factor (TNF), 102 Tyr, 70

U [URE3], 128 -helical model, 162 filaments, 145 functional domain of, and its conformation, 148 genetic selection criteria for, 131 nomenclature, 130 prion domains of, 129, 135–136 protease resistance, 145–146 STEM micrographs, 155 superpleated -structure model for, 157–159 [ure-o] cells, 128 Ure2p fibrils, structural model for, 260–262 Ure2p protein, 128. See also [URE3] aggregation of, 138 filament polymorphism in, 164–165 filaments, 142 filaments, electron diffraction of, 147 filaments formation and conformations, 149, 151 unfolded conformations, 147

320

SUBJECT INDEX

V Valine, 74, 195 Varied morphology, of amyloid fibrils, 267 Viral fibers, crystal structures of adenovirus, reovirus, and phage PRD1 fiber head domains, 99–102 adenovirus, reovirus, and phage PRD1 fiber shaft domains, 102–107 morphology, 99 tailed phage fibers, 107–111 Viral fibrous proteins, 111 Viru fibers, 97, 99 Virulence factors abundance of, 56 bacterial, 67, 71 of Gram-negative bacteria, 83 resident, 85

W Water-filled nanotube model for amyloid fibrils, 60, 204–205. See also Cylindrical -helix model Water spinning, 21

Wild-type PABPN1 (N-WT), fibrils from N-terminal domain of, 202

X X-ray crystallography, 182 X-ray diffraction study amyloidogenic core domains, 198–199 polyalanine, 199–201 of prion peptides, 191–192, 194 X-ray fiber diffraction, 184 X-ray study, for PrP peptides, 196

Y YadA adhesin, 76 Yeast cells, wild-type, 128 prion fibrils, polymorphism of, 12 prion filaments, models of, 151–156 Yeast Ure2p prion, 116 Yersinia enterocolitica, 88

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