Advances in Protein Chemistry, Volume 57: Prion Proteins (Advances in Protein Chemistry)

ADVANCES IN PROTEIN CHEMISTRY Volume 57 Prion Proteins

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ADVANCES IN PROTEIN CHEMISTRY EDITED BY FREDERIC M. RICHARDS

DAVID S. EISENBERG

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

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

VOLUME 57

Prion Proteins EDITED BY BYRON CAUGHEY National Institutes of Health, Rocky Mountain Laboratories, Hamiliton, Montana

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CONTENTS

PREFACE

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Prion Protein Diversity and Disease in the Transmissible Spongiform Encephalopathies SUZETTE A. PRIOLA I. II. III. IV. V. VI. VII.

General Background . . . . . . . . . . . . . . . PrP Biosynthesis . . . . . . . . . . . . . . . . . . Conversion of PrP-sen into PrP-res. . . . . . . . PrP and TSE Strains . . . . . . . . . . . . . . . . PrP-res Formation and the TSE Species Barriers PrP-res and Familial TSE . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . .

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Mass Spectrometric Analysis of Prion Proteins MICHAEL A. BALDWIN I. Modern Mass Spectrometric Techniques for Protein Characterization . . . . . . . . . . . . . . . . . . . . . II. Identification and Preliminary Analysis of PrP . . . . III. Confirmation of the PrPSc Amino Acid Sequence . . IV. Non-PrP Peptides in Prion Preparations. . . . . . . . V. N -Linked Oligosaccharides . . . . . . . . . . . . . . . VI. Analysis of the GPI Anchor . . . . . . . . . . . . . . . VII. Analysis of Intact PrP by MALDIMS and ESIMS . . . v

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VIII. IX. X. XI.

Processing of Chicken PrP . . . . . . . . . Recombinant PrP and Synthetic Peptides . Accessory Molecules in Scrapie Prions. . . Conclusions . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . .

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I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. The NMR Structures of the Recombinant Bovine, Human, Mouse, and Syrian Hamster Prion Proteins . . . . . . . . . . . . . . . . . . III. Prion Protein Structure and the Species Barrier . . . . . . . . . . . IV. Conclusions and Outlook . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Three-Dimensional Structures of Prion Proteins KURT WÜTHRICH AND ROLAND RIEK

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Folding Dynamics and Energetics of Recombinant Prion Proteins RUDI GLOCKSHUBER I. II. III. IV.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Folding of Recombinant PrPC . . . . . . . . . . . . . . . . . . . . . The Role of the Single Disulfide Bond of PrP. . . . . . . . . . . . . Influence of Point Mutations Linked with Inherited Human Prion Diseases on the Thermodynamic Stability of Recombinant PrPC . . V. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Simulations and Computational Analyses of Prion Protein Conformations DARWIN O.V. ALONSO AND VALERIE DAGGETT I. PrP Conformational Transitions . . . . . . . . . . . . . . . . . . II. Predictions of PrP Structures and Studies of Peptide Fragments to Test the Models . . . . . . . . . . . . . . . . . . . . . . . . . . III. PrPC Structural Models from NMR. . . . . . . . . . . . . . . . . IV. Detailed Modeling Studies Based on NMR Models . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Interactions and Conversions of Prion Protein Isoforms BYRON CAUGHEY, GREGORY J. RAYMOND, MICHAEL A. CALLAHAN, CAI’NE WONG, GERALD S. BARON, AND LIANG-WEN XIONG I. II. III. IV. V.

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I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. The Amyloid Peptides of Gerstmann-Stäussler-Scheinker Disease . III. Unraveling the Conformational Conversion of PrPC to PrPSc Using Synthetic Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Unraveling the Pathogenesis of Prion Diseases Using Synthetic Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . TSE-Associated Changes in PrP . . . . . . . . . . . . . . . . . . . Mechanistic Models of PrP-res Formation . . . . . . . . . . . . . . Binding Interactions between PrP-sen and PrP-res . . . . . . . . . PrPSc-Induced Conversion of PrP-sen and PrP-res: Biological Connections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mechanistic Studies of the PrP-Res-Induced Conversion Reaction PrP-sen/PrP-res Interactions and Species Barriers . . . . . . . . . TSE Studies and PrP-sen/PrP-res Interactions . . . . . . . . . . . PrP Perturbations and TSE Infectivity . . . . . . . . . . . . . . . . PrP-sen/PrP-res Interactions and the Search for Anti-TsE Drugs . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Studies of Peptide Fragments of Prion Proteins FABRIZIO TAGLIAVINI, GIANLUIGI FORLONI, PASQUALINA D’URSI, ORSO BUGIANI, AND MARIO SALMONA

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Biosynthesis and Cellular Processing of the Prion Protein DAVID A. HARRIS I. II. III. IV.

Introduction . . . . . Cell Biology of PrPC . Cell Biology of PrPSc . Conclusions . . . . . References . . . . . .

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Interaction of Prion Proteins with Cell Surface Receptors, Molecular Chaperones, and Other Molecules SABINE GAUCZYNSKI, CHRISTOPH HUNDT, CHRISTOPH LEUCHT, AND STEFAN WEISS I. II. III. IV. V.

Introduction . . . . . . . . . . . . . . Cell Surface Receptors . . . . . . . . Molecular Chaperones of Mammals . Interaction between Prion Proteins . Other PrP Interacting Molecules. . . References . . . . . . . . . . . . . . .

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Transgenic Studies of the Influence of the PrP Structure on TSE Diseases EMMANUEL A. ASANTE AND JOHN COLLINGE I. II. III. IV. V. VI. VII. VIII. IX. X. XI.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . Effects of PrP Gene Ablation . . . . . . . . . . . . . . . . . PrPC Is Necessary for Disease Propagation . . . . . . . . . Structure and Function of the PrP Gene . . . . . . . . . . Transgenic Studies of PrP Topology . . . . . . . . . . . . . Spontaneous Disease in Mutant Transgenic Mice . . . . . Transgenic Studies of the Species Barrier . . . . . . . . . . Transgenic Studies of Incubation Period . . . . . . . . . . Transgenic Studies of the Molecular Basis of Prion Strains Transgene Vector Considerations . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Yeast Prions Act as Genes Composed of Self-Propagating Protein Amyloids REED B. WICKNER, KIMBERLY L. TAYLOR, HERMAN K. EDSKES, MARIE-LISE MADDELEIN, HIROMITSU MORIYAMA, AND B. TIBOR ROBERTS I. II. III. IV. V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genetic Criteria for Yeast Prions . . . . . . . . . . . . . . . . . . [URE3] and URE2 Affect Nitrogen Catabolite Repression. . . . [PSI] and SUP35 Affect Efficiency of Translation Termination . [URE3] and [PSI] as Prions of Ure2p and Sup35p, Respectively

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VI. The Prion Domains of Ure2p. . . . . . . . . . . . . . . . . . . . . VII. Further Genetic Evidence That [URE3] Is a Prion . . . . . . . . . VIII. Ure2p Is Protease Resistant in Extracts and Aggregated in Vivo in [URE3] Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX. Amyloid Formation in Vitro by Ure2p . . . . . . . . . . . . . . . . X. [Het-s], a Prion of the Fungus Podospora anserina, Is Necessary for a Normal Function . . . . . . . . . . . . . . . . . . . . . . . . XI. Comparison of the Evidence for Yeast Prions with That for TSEs XII. Implications of Yeast-Prion Amyloidoses and the Podospora Prion . XIII. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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[PSI+], SUP35, and Chaperones TRICIA R. SERIO AND SUSAN L. LINDQUIST I. II. III. IV. V. VI. VII. VIII.

[PSI+] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Formulation of the Prion Hypothesis . . . . . . . . . . . . . . . Genetic and Cell Biological Support for [PSI+] as a Yeast Prion . A Model for the [PSI+] Phenotype . . . . . . . . . . . . . . . . . Crucial Residues in the Replication of Protein States . . . . . . Modeling [PSI+] in Vitro . . . . . . . . . . . . . . . . . . . . . . . Regulation of [PSI] Metabolism by Molecular Chaperones . . . Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

AUTHOR INDEX SUBJECT INDEX

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PREFACE

The epidemic of bovine spongiform encephalopathy (BSE or mad cow disease) in Europe and its recent spread to humans have sharpened interest in the molecular processes that underlie the transmissible spongiform encephalopathies (TSE) or prion diseases. Research in this area has expanded markedly over the last several years leading to dramatic advances in deciphering the chemistry of prion proteins (PrPs). High-resolution NMR structures of several recombinant PrP molecules have been solved. Folding, unfolding, and aggregation pathways have been characterized. Interactions and interconversions between normal and TSE-associated forms of PrP have been explored. PrP ligands have been identified and possible physiological roles suggested. These new insights have suggested ways in which misfolding and/or polymerization of PrP could be critical in TSE diseases of mammals. A predominant and long-standing hypothesis is that an abnormal pathologic form of PrP, e.g. PrPSc, propagates itself as an infectious protein-based, nucleic acid-free pathogen, or prion, by inducing the conversion of the host’s normal PrP to the pathologic form. Although some unsettling gaps remain in the proof of the prion hypothesis for TSE diseases, prion-like phenomena have been convincingly demonstrated in yeast and other fungi. Fungal prions are more amenable to clear genetic, biochemical, and mechanistic characterization and provide solid “proofs of principle” that prions exist in biology as protein-based elements of inheritance and/or pathogens. At the molecular level, the most fundamental process in these phenomena appears to be self-propagating changes in protein folding and polymerization. This volume of Advances in Protein Chemistry provides detailed and authoritative reviews of the current approaches to understanding the various prion-related proteins and the challenges that remain in xi

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describing the molecular bases for TSE diseases and other prion-like phenomena. It should prove useful both to members of the TSE and fungal prion fields and to those outside the field who seek a comprehensive understanding of these topics from the point of view of protein structure. Prion protein nomenclature or “Abandon hope all ye who enter here”: There is no doubt that the nomenclature of the TSE field can be confusing to outside observers and seasoned researchers alike. Part of this confusion arises from the genuine uncertainty that exists about the normal function(s) of PrP, the nature of TSE infectivity, and the various and variable abnormal properties of PrP most relevant to infectivity and/or TSE pathogenesis. No attempt has been made to unify the terminology used in the different chapters of this volume for two reasons. First, this seems like a hopeless task. Second, it is my belief that the nomenclature in any rapidly changing field should evolve with the facts as they are revealed. This state of flux will naturally require that authors carefully define and redefine their usage of terminology from time to time. Unfortunately, it also may require that readers of literature have the patience of Job and the semantic acumen of Supreme Court justices. Nonetheless, I can offer a few generalities that may aid the uninitiated reader at the outset. When used alone, the term “prion” refers to the infectious agent and has most recently been defined by its originator, Stanley Prusiner, as “proteinaceous infectious particles that lack nucleic acid” [Proc. Natl. Acad. Sci. USA 95:13363-13383 (1998)]. Thus the use of the term tends to imply a belief that the agent lacks nucleic acid. Those who are fully convinced that this belief is correct use the term “prion” without batting an eyelash. Those who remain unconvinced or uncertain stick to the generic term “infectious agent” or are content to use “prion” as if it means “that weird infectious TSE agent, whatever it turns out to be.” The combined term “prion protein” (PrP) refers to the protein rather than the infectious agent. However, it is increasingly apparent that PrP exists in a wide variety of forms that are normal (e.g., PrPC), TSE-associated (e.g., PrPSc, PrPCJD, PrPBSE), neither (e.g., various recombinant and/or mutated versions of the PrP structure), or both. This structural diversity complicates the use of Prusiner’s original terms PrPC (cellular PrP) and PrPSc (scrapie PrP). These terms are still useful under many circumstances, but often need further, more operational and/or functional qualification as the sophistication of current studies increases. For instance, since protease resistance is most commonly used to discriminate normal and TSE-associated forms of PrP, the operational terms PrP-sen (protease-sensitive PrP) and PrP-res (protease-

PREFACE

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resistant PrP) were initiated [J. Virol. 64:1093–1101 (1990)]. However, even the use of the terms PrP-sen and PrP-res is now complicated by the realization that PrP molecules can exhibit qualitatively and quantitatively different states of sensitivity or resistance to proteolysis. Ultimately, one would hope the naming of various PrP structures will be aided by continuing advances in our understanding of the functions of these fascinating and enigmatic molecules. Byron Caughey

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PRION PROTEIN DIVERSITY AND DISEASE IN THE TRANSMISSIBLE SPONGIFORM ENCEPHALOPATHIES BY SUZETTE A. PRIOLA Laboratory of Persistent Viral Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Rocky Mountain Laboratories, Hamilton, Montana 59840

I. General Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. TSE Disease and Prion Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. PrP Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. PrP-sen. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. PrP-res . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Conversion of PrP-sen into PrP-res . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Models of PrP-res Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. PrP-res “Replication” . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Role of Cofactors in the Conversion of PrP-sen to PrP-res . . . . . . . . . . . . D. Co- or Posttranslational Modifications of PrP and Disease . . . . . . . . . . . . IV. PrP and TSE Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Strains and Protein Conformation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Strains and PrP Glycosylation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Strains and the Host . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. PrP-res Formation and the TSE Species Barriers . . . . . . . . . . . . . . . . . . . . . . . A. TSE Species Barriers and PrP Primary Sequence . . . . . . . . . . . . . . . . . . . . B. Species-Specific Formation of PrP-res . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. PrP Sequence and Disease Susceptibility . . . . . . . . . . . . . . . . . . . . . . . . . . VI. PrP-res and Familial TSE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Spontaneous Formation of PrP-res . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Mechanisms of Disease in Familial TSE Diseases . . . . . . . . . . . . . . . . . . . . VII. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. GENERAL BACKGROUND A. Introduction In the late 1970s, Great Britain altered the process by which animal carcasses were rendered to provide meat and bone meal protein supplements to sheep, cattle, and other livestock. Several years later a new disease was recognized in the British cattle population. The pathological and immunohistochemical characteristics of the disease placed it among a group of diseases known as the transmissible spongiform encephalopathies (TSE), and the new syndrome was named bovine spongiform encephalopathy (BSE) (Wells et al., 1987; Bradley and Wilesmith, 1993; Anderson et al., 1996). Despite concerns that the human 1

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population might be exposed to BSE there was no evidence that humans were susceptible to scrapie, a sheep TSE that had been recognized in England for over 200 years. This suggested that humans might not be susceptible to BSE. However, approximately 10 years after BSE was first recognized, a previously unknown form of the human TSE, Creutzfeldt-Jakob disease (CJD), was identified in young people in Great Britain (Will et al., 1996). The suggestion that this new type of CJD was the consequence of exposure of humans to BSE has now been supported by several different studies (Collinge et al., 1995; Collinge et al., 1996; Bruce et al., 1997; Raymond et al., 1997). More than 90 cases of variant CJD (vCJD) have now been confirmed. Thus, vCJD represents the emergence of the TSE as a potentially widespread health threat to the human population and reinforces the need to determine the mechanisms underlying TSE pathogenesis. B. TSE Disease and Prion Protein Transmissible spongiform encephalopathies are a group of rare, fatal, and transmissible neurodegenerative diseases, which, in addition to scrapie and BSE, include chronic wasting disease (CWD) in mule deer (Williams and Young, 1980) and elk (Williams and Young, 1982), and transmissible mink encephalopathy (TME) (Marsh and Kimberlin, 1975). TSE diseases are characterized by accumulation, primarily in the brain and lymphoreticular system, of an abnormal form of the normal host-encoded prion protein (PrP). Normally PrP, an approximately 254 amino acid glycoprotein that is expressed on the cell surface in a wide variety of tissues, is both soluble and sensitive to digestion with proteinase-K (PrP-sen).1 By contrast, the abnormal form of PrP associated with TSE disease (PrP-res) forms insoluble aggregates and is partially resistant to digestion with proteinase K. In PrP-res, only the N-terminal 60 or so amino acids are cleaved. As a result, PrP- res migrates 6 to 7 kDa lower than PrP-sen on SDS-PAGE gels (Fig. 1). In the brain, PrP-res can demonstrate the properties of classic amyloid and form β-sheet rich, insoluble protein deposits that exhibit birefringence under crosspolarized light after staining with the amyloid dye Congo red. In some forms of the disease, PrP-res is also associated with amyloid plaques. Thus, the TSE diseases are often considered another type of amyloid 1 The normal form of PrP has been referred to as either PrPC for PrP-cellular or PrP-sen, which designates its sensitivity to proteinase K. The abnormal form of PrP has been referred to as either PrPSc for PrP-scrapie or PrP-res, which designates its partial resistance to proteinase K. Since the terms PrP-sen and PrP-res reflect a general biochemical definition rather than a disease-specific phenotype, they will be used throughout this discussion.

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FIG. 1. Abnormal PrP is partially resistant to proteinase K. PrP-sen and PrP-res from (A) hamster brains infected with the hamster scrapie strain 263K, (B) cell-free conversion assay (Kocisko et al., 1994) using radiolabeled hamster PrP-sen and hamster PrP-res isolated from infected hamster brains, and (C) Mouse PrP-sen and PrP-res derived from mouse scrapie-infected murine neuroblastoma cells. In the absence of proteinase K (PK), PrP-sen migrates as three major bands representing the 25 kDa unglycosylated (U), 30 kDa partially glycosylated (P), and ~32–40 kDa fully glycoslyated forms (F). These bands would disappear if the protein were treated with PK (not shown). PrP-res is partially resistant to PK and only the N-terminal 60 or so amino acids are cleaved. The resultant unglycosylated, partially glycosylated and fully glycoslylated bands therefore migrate ~6–7 kDa lower on the gel. *, oligomeric PrP. For each panel, molecular mass markers in kilodaltons are shown on the right.

disease such as Alzheimer’s disease, Huntington’s disease, or type II diabetes (Chesebro and Caughey, 1997). The TSEs are unique among the amyloid diseases in that they are transmissible. However, no virus or bacteria has ever been associated with TSE infectivity, suggesting the possibility of a unique disease etiology. The extreme resistance of TSE infectivity to inactivation by a variety of harsh treatments led to the hypothesis that the infectious agent in these diseases was an infectious, self-replicating protein (Alper et al., 1967; Griffith, 1967). The fact that PrP-res is found in TSE-infected tissues and is very closely associated with tissue preparations enriched for infectivity has led to the proposal that PrP-res itself is the infectious protein agent (Prusiner, 1982). Although this hypothesis remains a point of contention in TSE research, the key role of PrP in TSE pathogenesis is undeniable (Bueler et al., 1993; Brandner et al., 1996; Blattler et al., 1997) and understanding the manner in which this protein influences disease is critical to understanding the molecular mechanisms behind the TSE diseases.

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II. PrP BIOSYNTHESIS A. PrP-sen The biosynthesis of PrP-sen and PrP-res has been studied using normal mouse neuroblastoma (MNB) cells, which only express PrP-sen, and mouse scrapie-infected neuroblastoma (Sc+-MNB) cells, which are persistently infected with mouse scrapie and express both PrP-sen and PrP-res (Fig. 1C). The synthesis of PrP-sen in uninfected and scrapieinfected MNB cells appears to be the same, but in Sc+-MNB, where PrPsen is the precursor to PrP-res (Borchelt et al., 1990; Caughey and Raymond, 1991), the biosynthesis of PrP-res differs dramatically from that of PrP-sen (Caughey et al., 1989; Borchelt et al., 1990; Caughey and Raymond, 1991; Caughey et al., 1991a; Taraboulos et al., 1992). PrP-sen is transferred cotranslationally into the endoplasmic reticulum where the N-terminal signal peptide is cleaved and cleavage of the C-terminus is followed by addition of a glycophosphotidylinositol (GPI) membrane anchor (Fig. 2A) (Hope et al., 1986; Bolton et al., 1987; Hope et al., 1988). Following the addition of high mannose glycans to one or two potential N-linked glycosylation sites (Fig. 2A), PrP is translocated into the Golgi apparatus where the high mannose glycans are converted to complex or hybrid glycans (Caughey et al., 1989). The most common form of PrP-sen therefore migrates as three discrete bands consisting of unglycosylated, partially glycosylated and fully glycosylated PrP (Fig. 1). After translocation to the cell surface, PrP is anchored to the plasma membrane via the GPI anchor (Stahl et al., 1987; Caughey et al., 1989; Stahl et al., 1990), and the majority of the cell surface PrP is phospholipase sensitive (Caughey et al., 1989; Caughey et al., 1990). Once on the cell surface, PrP has a half-life of 3 to 6 hours (Caughey and Raymond, 1991). Over time, most PrP appears to be degraded in a nonacidic compartment bound by cholesterol-rich membranes (Taraboulos et al., 1995), although some is released into the tissue culture medium (Caughey et al., 1988; Caughey et al., 1989; Borchelt et al., 1990; Caughey and Raymond, 1991). B. PrP-res Both the normal and abnormal forms of PrP are derived from the same gene and have the same amino acid sequence. Pulse-chase studies in Sc+MNB have demonstrated that PrP-res is in fact posttranslationally derived from mature PrP-sen (Borchelt et al., 1990; Caughey and Raymond, 1991). Unlike PrP-sen, PrP-res is resistant to phospholipases and protease treat-

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FIG. 2. Conversion of PrP-sen into PrP-res. (A) Unprocessed PrP-sen from amino acids 1–254 is shown. The scissors pointing down at the N-terminus indicate the signal peptide cleavage site while the scissors pointing up at the C-terminus indicate the GPI anchor addition cleavage site. The GPI anchor is indicated by the lollipop at the C-terminus. The two N-linked glycosylation sites are indiated by the branched vertical lines. Open squares represent the octapeptide repeat region where each square equals one repeat. The areas of secondary structure based on hamster PrP-sen NMR data (Liu et al., 1999) are indicated by the black and gray squares where S, β strand, H, α helix. The line designates areas of loop or disordered structure. The asterisks show the region where PK can digest PrP-res to give the truncated form of PrP-res shown in (B). The structure shown for hamster PrP-res is based solely on predictions from infrared spectroscopy measurements using the method of Garnier (Caughey et al., 1991b) and thus remains hypothetical. The dashed portion of the line indicates the variability of the C-terminus of PrP-res after PK digestion. Note the higher beta-sheet content and the fact that most of the predicted beta-sheet structures in PrP-res are downstream of the S1 and S2 betastrand regions of PrP-sen. These strands have been hypothesized to form a potential “nucleation site” that triggers the formation of PrP-res (Riek et al., 1996).

ments (Borchelt et al., 1990; Caughey et al., 1990; Stahl et al., 1990; Safar et al., 1991) and appears to accumulate intracellularly in Sc+-MNB cells with little, if any, expressed on the cell surface (Borchelt et al., 1990; Caughey et al., 1990; McKinley et al., 1991). The conversion of PrP from the proteasesensitive to protease-resistant form likely occurs either on the plasma membrane or along an endocytic pathway (Caughey and Raymond, 1991;

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Caughey et al., 1991a; McKinley et al., 1991), possibly in a cellular compartment containing cholesterol-rich membranes (Taraboulos et al., 1995). In Sc+-MNB cells, PrP-res eventually accumulates in the secondary lysosomes or on the cell surface (Caughey and Raymond, 1991; Caughey et al., 1991a; McKinley et al., 1991). In vivo, it can also accumulate in the extracellular spaces as amorphous deposits or more organized fibrils or amyloid plaques (Jeffrey et al., 1994a; Jeffrey et al., 1994b). Despite its exposure to endolysosomal hydrolases, PrP-res has a half-life of more than 48 hours (Borchelt et al., 1990; Caughey and Raymond, 1991). The metabolic stability of PrP-res could explain its accumulation in vivo, especially in the nondividing cells of the central nervous system. III. CONVERSION OF PrP-SEN INTO PrP-RES There are no posttranslational modifications that can account for the different biochemical properties of PrP-sen and PrP-res, and it is likely that the difference between the two molecules is a conformational one (Fig. 2). Analysis of PrP-res by infrared spectroscopy and circular dichroism (Caughey et al., 1991b; Pan et al., 1993; Safar et al., 1993a; Caughey et al., 1998) and the recent determination of the structure of several different species of PrP-sen by nuclear magnetic resonance (NMR) (Fig. 3 and Wüthrich, this volume) (Riek et al., 1996; Donne et al., 1997; Riek et al., 1997; Liu et al., 1999; Garcia et al., 2000; Zahn et al., 2000) support this hypothesis and demonstrate that PrP-res has a much higher β-sheet content than PrP-sen (Fig. 2B). Thus, a key question within the TSE is how exactly the conformational conversion of PrP-sen to PrP-res occurs. This process is critical to disease pathogenesis, especially if PrP-res is the infectious agent, and any model of PrP-res formation must be able to explain at a protein level how PrP-res could (1) induce its own formation (i.e., replicate), (2) account for different scrapie strains in animals with one PrP genotype, (3) control not only disease incubation time but also species barriers to infection and, finally, (4) lead to some forms of human TSE that are heritable. In vivo studies in normal and transgenic mice, tissue culture studies in Sc+-MNB cells, and in vitro studies utilizing a recently developed cell-free conversion assay system have led to valuable new insights into PrP-res formation and its influence on the disease process. A. Models of PrP-res Formation Two primary models have been used to describe the self-propagation of PrP-res. The heterodimer model (Griffith, 1967; Bolton and Bend-

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FIG. 3. NMR structures of PrP-sen. The refined NMR structure of (A) recombinant mouse PrP-sen from amino acid residues 121–231 (Riek et al., 1996) and (B) recombinant hamster PrP-sen from amino acid residues 23–231 (Liu et al., 1999). The structure of full-length mouse PrP-sen has been derived (Riek et al., 1997), but residues from the signal peptide cleavage site to the structure illustrated (i.e., 23–120) exist in a flexible, “random coil-like” state as shown for the hamster PrP-sen structure in (B). S, β strand; H, α helix. Figure prepared with the program MOLMOL (Koradi et al., 1996).

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heim, 1988; Prusiner, 1991) proposes that a monomer of PrP-res interacts with a monomer of PrP-sen and converts it to more PrP-res by inducing a conformational change (see Caughey, this volume). The two molecules, now both PrP-res, separate and the process repeats. In the seeded polymerization model (Gadjusek, 1988; Jarrett and Lansbury, Jr., 1993), a “seed” composed of an organized array of multiple PrP-res molecules interacts with PrP-sen and converts it to PrP-res by inducing a conformational change. The newly formed PrP-res molecule remains a part of the polymer and the process continues, eventually leading to accumulation of PrP-res and disease. Most of the currently available evidence supports the seeded polymerization model. In yeast, the only other organism where prion-like activity has been found (Wickner, 1994), only aggregated forms of the yeast prion protein can induce the conversion (Glover et al., 1997; Paushkin et al., 1997; Taylor et al., 1997). Similarly, recent data suggest that only aggregates of PrP-res can induce the conversion of PrP-sen to PrP-res (Caughey et al., 1997). Dissociation of PrP-res to a soluble form leads to a loss not only of its “converting activity” but also of infectivity (Gasset et al., 1993; Safar et al., 1993b; Caughey et al., 1997; Shaked et al., 1999). Finally, monomeric PrP-res as resistant to proteinase K as aggregated PrP-res has never been detected, although it is still possible that less proteinase K-resistant monomeric PrP-res is present and active at low levels. B. PrP-res “Replication” The most direct evidence that PrP-res can induce its own formation comes from an experimental system in which radiolabeled tissue culture derived PrP-sen mixed with PrP-res isolated from TSE-infected brains can be converted into a proteinase K-resistant form (Kocisko et al., 1994; Kocisko et al., 1995; Kocisko et al., 1996). Via an interaction that may involve two separate domains in PrP (see Caughey, this volume), PrP-res binds selectively to PrP-sen and conversion of PrP-sen to a protease-resistant form occurs (Kocisko et al., 1996; Bessen et al., 1997; DebBurman et al., 1997; Horiuchi et al., 1999). In terms of size and glycoform ratio, the protease-resistant PrP formed in vitro is virtually indistinguishable from PrP-res isolated from infected brain (compare Fig. 1A and B). Some evidence suggests that binding and conversion are two distinct events (DebBurman et al., 1997), a hypothesis that is supported by data suggesting that initial PrP-PrP interactions may map to different regions of PrP than those that appear to influence conversion (Scott et al., 1992;

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Kocisko et al., 1995; Priola and Chesebro, 1995; Chabry et al., 1998; Chabry et al., 1999; Horiuchi et al., 1999). That PrP-res can induce PrPsen to convert to more of the abnormal form in a cell-free environment provides compelling evidence that PrP-res can “replicate” itself in the absence of any metabolic activity and demonstrates that a viable virus or bacteria is not needed for this process to occur. C. Role of Cofactors in the Conversion of PrP-sen to PrP-res The role of other molecules in the conversion process is unclear (Weiss, this volume). Glycosaminoglycans (GAGs) have been proposed as one possible cofactor (see Priola and Caughey, 1994 for review). GAGs bind PrP (Gabizon et al., 1993; Caughey et al., 1994), inhibit formation of PrP-res (Caughey and Raymond, 1993), are often associated with PrP plaques in the brain of infected individuals (Snow et al., 1989), and can inhibit TSE disease in vivo (Kimberlin and Walker, 1986). Under certain conditions, GAGs may even enhance formation of protease-resistant PrP (Wong et al., 2001). Chaperones can enhance the formation of PrP-res in the cell-free conversion system, although none of the chaperones tested are found in cellular compartments where PrP-res formation most likely occurs (DebBurman et al., 1997). Other as yet unidentified cell factors have also been suggested as potential cofactors in the conversion process (Telling et al., 1995; Kaneko et al., 1997). It is certainly possible that any or all of these proposed cofactors could play a role in formation of PrP-res. D. Cotranslational or Posttranslational Modifications of PrP and Disease 1. The Disulfide Bond Both PrP-sen and PrP-res contain a single intramolecular disulfide bond (Fig. 2) (Turk et al., 1988). The importance of disulfide bonds in TSE disease was first suggested by early experiments demonstrating that treatment of mouse scrapie-infected brain fractions with SDS and 2mercaptoethanol significantly reduced scrapie infectivity (Somerville et al., 1980). Consistent with these in vivo results, the presence of the disulfide bond appears to affect PrP folding and PrP-res formation. Removal of the disulfide bond in PrP-sen leads to a protease-resistant form of PrP (Jackson et al., 1999; Ma and Lindquist, 1999) and prevents its conversion to protease resistance in an in vitro conversion assay (Herrmann and Caughey, 1998).

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2. Glycosylation and the GPI Anchor PrP-sen is glycosylated and contains a GPI anchor, both of which could effect formation of PrP-res. While the GPI membrane anchor does not appear to be essential, recent evidence shows that the manner in which PrP-sen is inserted in the cellular membrane can influence its biochemical properties as well as disease pathogenesis (Lehmann and Harris, 1996; Kaneko et al., 1997; Hegde et al., 1998). Both deglycosylated PrP-sen and GPI anchor minus PrP-sen are efficiently converted to PrP-res in vitro (Taraboulos et al., 1990; Rogers et al., 1993; Kocisko et al., 1994). However, removal of one or both of the PrP glycosylation sites changes the biochemical processing of PrP-sen and can alter TSE pathogenesis in transgenic mice depending on the strain of infectious agent (DeArmond et al., 1997). Thus, while N-linked glycosylation and the GPI anchor are not essential for PrP-res formation in vitro, in vivo glycosylation of PrP-sen may influence certain aspects of TSE disease. 3. Metal Binding Metals may also be involved in PrP function and/or formation of abnormal PrP. Peptides encompassing the N-terminal octapeptide motif of PrP have been shown to bind copper (Hornshaw et al., 1995a; Hornshaw et al., 1995b). Binding of copper Cu(II) can change the structural properties of the peptide resulting in a more ordered, helical conformation (Miura et al., 1996; Smith et al., 1997). Thus, divalent metal cations may be important in determining PrP structure at the flexible N-terminus (Fig. 3B). Full PrP-sen has also been shown to bind Cu(II) (Brown et al., 1997a; Stockel et al., 1998) and act as a superoxide dismutase (Brown and Besinger, 1998; Brown et al., 1999), implicating its possible involvement in mechanisms of oxidative stress (Brown et al., 1997a; Brown et al., 1997b; Giese et al., 1998). Finally, PrP-sen, which has bound manganese Mn(II) instead of Cu(II), becomes more proteaseresistant over time and loses any superoxide dismutase activity (Brown et al., 2000). Cumulatively these studies suggest the possibility that divalent cation binding to PrP-sen, resulting in a loss of PrP-sen function and/or accumulation of protease-resistant PrP, could be involved in the neurodegenerative processes of the TSE diseases. IV. PrP-RES AND TSE STRAINS Strains in the TSE are defined by differences in brain pathology, disease incubation time, clinical disease, and species tropism (for review see Bruce, 1996). Strains of TSE agent have been described in humans

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(Parchi et al., 1996), sheep (Kascsak et al., 1985; Kascsak et al., 1986), and hamsters (Bessen and Marsh, 1994). At least 20 strains have been extensively characterized and studied in experimental mouse models of scrapie (Bruce, 1996). It is especially challenging to explain TSE strains in terms of PrP-res formation because multiple strains have been described in species with one PrP genotype. Because the TSE appear to be a disease of protein folding, the answer may reside not in the PrP amino acid sequence but rather in the conformation of the protein. A. Strains and Protein Conformation If PrP-res can adopt several stable variations in structure, it might be able to pass on its own strain-specific characteristics to PrP-sen via direct PrP-PrP interactions. Support for this hypothesis was first derived from studies of hamster-adapted TME strains (Bessen and Marsh, 1992b; Bessen and Marsh, 1994). As a result of a difference in the PK-cleavage site in the N-terminus of PrP-res, protease digestion of the hyper and drowsy hamster TME strains yielded two different sized protease-resistant products (Bessen and Marsh, 1994). This strongly suggested that the PrP-res isolated from hyper or drowsy infected hamster brain had different protein conformations (Bessen and Marsh, 1994). Later studies using transgenic mice infected with different strains of human CJD also implied that TSE strains could be dependent on the conformation of PrP-res (Collinge et al., 1996; Telling et al., 1996b). In vitro, there is direct evidence that PrP-res from animals infected with different strains of TSE agent can faithfully propagate strain-specific protein conformations (Bessen et al., 1995; Bessen et al., 1997). PrP-res isolated from hamsters infected with the hyper or drowsy hamster adapted TME strains can convert PrP-sen to protease-resistant forms, which differ in size by ~1 to 2 kDa depending on the strain used. This strain-specific conversion occurs even when the PrP-res has been pretreated with proteinase K to remove other potential contaminating proteins (Fig. 4). Therefore, PrP-res alone can determine the final conformation of the protease-resistant product (Bessen et al., 1995). Similarly, PrP-res isolated from mice infected with different mouse scrapie strains can determine strain-specific glycoform patterns (S. Priola, unpublished data). Recent structural data using infrared spectroscopy (Caughey et al., 1998) or a conformation-dependent immunoassay (Safar et al., 1998) have now demonstrated that PrP-res from different mouse and hamster TSE strains (including hyper and drowsy) do indeed have different conformations. Thus, there is now ample evidence to support the hypothesis that self-propagation of PrP-res may proceed via distinc-

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FIG. 4. PrP-res isolated from hamsters infected with different scrapie strains can determine the final conformation of the protease-resistant product. Radiolabeled hamster PrP-sen without the GPI anchor was mixed with hamster PrP-res isolated from the brains of hamsters infected with either hyper (Hy) or drowsy (Dy) scrapie in a cellfree conversion assay (Kocisko et al., 1994). The PrP-res was either pre-treated (+) or not (–) with proteinase K to remove any contaminating proteins (Bessen et al., 1995). The left side panel shows the radiolabeled PrP-sen in the reaction after a 2-day incubation and before digestion with proteinase K (–PK) and represents 10% of the total reaction. The right side panel shows the radiolabeled, protease-resistant product remaining after digestion of the reaction with proteinase K (+PK) and represents the remaining 90% of the reaction. Note the characteristic size shift of ~1 to 2 kDa which is used to distinguish PrP-res from the hyper and drowsy scrapie strains (Bessen and Marsh, 1992a). There is no significant difference in the amount of protease-resistant product formed when PK pretreated or untreated PrP-res is used (53% vs. 48% for hyper PrP-res and 47% vs. 40% for drowsy PrP-res). Molecular mass markers are shown in kilodaltons on the right.

tive three-dimensional structures, which in turn could provide some molecular basis for scrapie strains. B. Strains and PrP Glycosylation Overall, the role of posttranslational modifications to PrP in scrapie strains is poorly understood. Although in vitro studies have demonstrated that deglycosylated PrP-sen is efficiently converted to PrP-res (Taraboulos et al., 1990; Rogers et al., 1993; Kocisko et al., 1994), it is possible that glycosylation of PrP is still involved in strain-specific differences in PrP-res. Abnormal PrP is heterogeneous not only in the location of proteolytic cleavage sites (Somerville and Ritchie, 1990; Bessen and Marsh, 1994; Parchi et al., 1996; Chen et al., 1997) but also in N-linked glycosylation (Somerville and Ritchie, 1990). Furthermore, different strains of TSE agent are associated with different glycoform ratios (Collinge et al., 1996;

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Parchi et al., 1997; Somerville et al., 1997; Kuczius et al., 1998). Precisely how these strain-specific glycoform ratios are maintained is currently an open question. For example, they may be the consequence of preferential conversion of PrP-sen glycoforms into PrP-res. In this instance, it is the strain-specific PrP-res that determines the final glycoform pattern (Somerville, 1999). Alternatively, if there are differences in the PK sensitivities of the different forms of glycosylated PrP-res, degradation of the newly formed PrP-res in endolysosomal compartments could account for the different banding patterns. In this latter case, the different glycoform ratios would be the result of events occurring after the formation of PrPres (DeArmond et al., 1999). Either of these possibilities could be modulated by the ability of glycosylation to affect protein conformation and stability (O’Conner and Imperiali, 1996; O’Conner and Imperiali, 1998), which in turn could influence PrP/PrP interactions and the rate of PrPres formation (DeArmond et al., 1997). C. Strains and the Host Of course, the infected host must also play a role in the molecular aspects of TSE strain pathogenesis (Somerville, 1999). Differences in PrP-sen glycosylation in different populations of neurons of the brain could target incoming infectivity to populations of neurons with specific PrP-sen glycoforms. If this were the case, PrP-res would be restricted by the available PrP-sen glycoforms in a particular cell. Recent studies have shown that the number and isoelectric points of PrP-sen charge isomers differ in different regions of the brain (DeArmond et al., 1999). These data support the hypothesis that strain-specific PrP-res glycoform patterns could be dictated by cell-specific differences in PrP-sen glycoforms (Somerville, 1999). Further support comes from in vivo studies utilizing transgenic mice overexpressing hamster PrP-sen mutated at the Nlinked glycosylation sites, which suggest that neuronal targeting of infectivity may be important (DeArmond et al., 1997). By contrast, there are also experiments demonstrating that strain-specific glycoform patterns do not significantly differ in PrP-res isolated from several different brain regions (Kuczius et al., 1998). Clearly, the questions surrounding the mechanisms behind the strain-specific properties of PrP-res have not been fully resolved. V. PrP-RES FORMATION AND TSE SPECIES BARRIERS Species barriers in the TSE are defined as the resistance to TSE disease by one species after infection with the TSE agent of another species.

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This resistance is reflected by extremely long incubation times on first passage of a TSE agent into a new host species. The barrier to infection on first exposure can be very strong and can lead to incubation times that exceed the lifetime of the host. However, as the agent is serially passaged through the new host, adaptation occurs and incubation times shorten until a stable incubation time is reached. The fact that PrP-res binds PrP-sen (DebBurman et al., 1997; Horiuchi et al., 1999) suggests that the PrP amino acid sequence may be involved in the conversion process and that homology between the two molecules is important if conversion is to occur. Homologous PrP molecules may form PrP-res efficiently, leading to a more rapid accumulation of PrP-res to pathogenic levels and thus to clinical disease. Conversely, heterologous PrP molecules might have a reduced potential to form PrP-res, thus slowing the disease process. However, PrP-sen is a highly conserved protein among all mammalian species (Wopfner et al., 1999). If the primary sequence of PrP controls species barriers in the TSE diseases at the protein level, generation of PrP-res must be exquisitely sensitive to minor changes in the amino acid sequence of PrP. A. TSE Species Barriers and PrP Primary Sequence The first evidence to suggest that the primary sequence of PrP could profoundly influence TSE species barriers came from studies utilizing the strong barrier to infection that exists between hamsters and mice. Although mice are fully susceptible to mouse scrapie, they are resistant to infection with a particular strain of hamster scrapie. Mice infected with hamster scrapie do not become clinically ill within the lifetime of the animal even though it appears that infectivity can be sequestered in these animals (Race and Chesebro, 1998). However, transgenic mice that overexpressed hamster PrP-sen were fully susceptible to hamster scrapie (Scott et al., 1989). Subsequent studies demonstrated that the hamster/mouse species barrier could be crossed even when hamster PrP-sen expression was restricted to neurons (Race et al., 1995) or astrocytes (Raeber et al., 1997), but not T-cells or hepatocytes (Raeber et al., 1999). Therefore, even though mouse and hamster PrP-sen are highly homologous and differ at only 16 amino acid positions (i.e., 94% homology), these differences are sufficient to significantly effect crossspecies transmission of TSE infectivity. B. Species-Specific Formation of PrP-res In vivo, transgenic mouse studies have mapped the region of PrP important in transmission of TSE infectivity across species barriers to an

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area encompassing the middle portion of PrP that is highly homologous among many different species (Fig. 5) (Scott et al., 1993; Telling et al., 1995; Wopfner et al., 1999). In both Sc+-MNB and cell-free systems, species-specific conversion of PrP-sen to PrP-res maps to the same region (Scott et al., 1992; Kocisko et al., 1995; Priola and Chesebro, 1995), but these studies have further demonstrated that there is in fact an exquisite sequence specificity in PrP-res formation. For example, in Sc+-MNB cells, a single amino acid residue mismatch at position 138 was sufficient to control the species-specific formation of mouse PrP-res (Fig. 5) (Priola and Chesebro, 1995). In the cell-free conversion system, a single mismatch at position 155 was sufficient to control the speciesspecific formation of protease-resistant hamster PrP (Fig. 5) (S. Priola, unpublished results) while a mismatch at residue 171 in sheep PrP influenced the species-specific formation of protease-resistant sheep PrP (Fig. 5) (Bossers et al., 1997; Raymond et al., 1997). These studies imply that PrP-sen and PrP-res that mismatch at a single critical amino acid residue could influence TSE species barriers by affecting the efficiency of PrP-res formation. Indeed, in vivo there is evidence for a single amino acid difference in the PrP molecules of two species affecting cross-species transmission of TSE infectivity. A mismatch at residue 142 in goat PrP, the same amino acid as residue 138, which has been linked with mouse PrP-res formation in Sc+-MNB cells (Priola and Chesebro, 1995), has been associated with the resistance of some goats to sheep scrapie and BSE (Goldmann et al., 1996). Similarly, residue 171 in sheep PrP has been associated with resistance to both sheep scrapie and BSE (Goldmann et al., 1994; Belt et al., 1995; Hunter et al., 1996). C. PrP Sequence and Disease Susceptibility Single amino acid changes have not only been associated with species barriers in TSE diseases but also with susceptibility to TSE diseases within a species as well. In mice, short and long scrapie incubation times are controlled by polymorphisms at two different amino acid residues in PrP (Westaway et al., 1987; Moore et al., 1998). In humans, it is well known that homozygosity at codon 129 is associated with a higher susceptibility to CJD (Palmer et al., 1991) and the same polymorphism may influence CWD pathogenesis in elk (Fig. 5). Resistance of sheep to scrapie has been correlated with heterozygosity at amino acid residue 171 (Goldmann et al., 1994; Bossers et al., 1997). Less subtle mutations in PrP-sen have also been correlated with PrP-res formation. Deletion of the N-terminus of PrP-sen or PrP-res (Fig. 4) up to position 88 (through the octapeptide repeat region, Fig. 1) do not

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FIG. 5. Region of PrP associated with TSE species barriers is highly conserved among different mammalian species. The structure of human PrP-sen (Zahn et al., 2000) is shown on the top line and the legend is the same as in Fig. 2A. The region of PrP, which is involved in cross-species transmission of TSE infectivity and in the species-specific formation of PrP-res, is expanded below. The amino acid sequence of PrP within this region is shown using the single letter code and using human (hu) PrP as the primary sequence reference. Boxed residues indicate amino acids that have been shown to interfere with PrP-res formation and that have been implicated in resistance to disease. The asterisk indicates that residue 129 in human PrP (and 132 in elk PrP) are polymorphic. The closed circle shows the residue associated with resistance to BSE in goats (Goldmann et al., 1996) and species-specific formation of PrP-res in Sc+MNB cells (Priola and Chesebro, 1995). The closed square indicates the residue associated with the speciesspecific formation of protease-resistant hamster PrP in vitro (S. Priola, unpublished observations), and the open triangle indicates a residue in sheep associated with resistance to sheep scrapie and BSE (Goldmann et al., 1994; Belt et al., 1995; Hunter et al., 1996). Abbreviations: bo, bovine; Md, mule deer; ov-AQ, scrapie susceptible sheep; ovAR, scrapie resistant sheep; ha, hamster; mo, mouse.

affect PrP-res formation (Rogers et al., 1993). In vivo studies have shown that transgenic mice expressing similar PrP-sen deletion mutants are susceptible to TSE disease, although PrP-res accumulates to lower levels than in normal mice (Fischer et al., 1996; Telling et al., 1997). However, removal of residues further into the flexible N-terminus yield a PrP-sen molecule that is not converted to PrP-res (Holscher et al., 1998;

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Zulianello et al., 2000). Thus, some portions of the flexible N-terminus are apparently important in the generation of PrP-res. Expression of different PrP-sen molecules can significantly affect disease. In transgenic mice that express both mouse PrP-sen and hamster PrP-sen, mouse scrapie disease incubation times increase as the expression level of heterologous hamster PrP-sen increases (Scott et al., 1989; Race et al., 1995; Raeber et al., 1997). Mice heterozygous for the PrP gene demonstrate a relationship between PrP expression level and scrapie incubation times (Bueler et al., 1994; Manson et al., 1994). Conversely, hamster PrP-expressing transgenic mice in which the mouse PrP gene has been ablated develop hamster scrapie more rapidly than hamster PrP transgenic mice that still express mouse PrP (Prusiner et al., 1990; Raeber et al., 1997). Overall, these in vivo observations indicate that the presence of heterologous PrP can interfere with disease and that this interference is dependent on expression level. Support for this hypothesis has come from studies in Sc+-MNB cells. Expression of a heterologous mouse PrPsen mutated at 1 or 2 residues significantly inhibited PrP-res formation from the wild-type endogenous mouse PrP-sen (Fig. 5) (Priola et al., 1994). Furthermore, this interference appeared to be dependent on the overall expression level of the mutant PrP-sen molecule (Priola, 1999). At the molecular level, cell-free conversion assays have clearly shown that interference is related to the ratio of heterologous PrP-sen molecules to PrP-res (Horiuchi et al., 2000). These same studies also suggested that the mechanism of interference was not dependent on PrP sequence-specific binding of PrP-sen to PrP-res but rather on blocking of some postbinding conversion step (Horiuchi and Caughey, 1999; Horiuchi et al., 2000). Thus, for TSE infection within a species as well as cross-species transmission of TSE, heterologous PrP molecules act as a type of host resistance factor, whereas homologous PrP molecules act as a type of susceptibility factor. VI. PrP-RES AND FAMILIAL TSE TSE agent infectivity is detectable in the brains of individuals afflicted with familial TSE, although there is no correlation with exposure to an infectious agent (for review see Pocchiari, 1994). PrP-res is also detectable in most familial forms of human TSE, although the size may vary (Kitamoto et al., 1991; Tagliavini et al., 1991; Kitamoto et al., 1993; Tagliavini et al., 1994; Chen et al., 1995; Piccardo et al., 1996). The question then is how exactly abnormal PrP is generated in familial TSE in the absence of

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exposure to PrP-res from an exogenous source. Because these diseases are heritable, genetics must play a role. Given the critical role that PrP has in infectious forms of TSE, the most likely genetic component is the PrP gene itself. Familial TSE diseases have been correlated with particular mutations in the PrP gene, and in most families association of the mutant PrP genotype with onset of disease (i.e., penetrance) is 100%. As seen in Fig. 6, these mutations occur throughout most of the PrP molecule, although many are clustered toward the C-terminus. A. Spontaneous Formation of PrP-res Although there is good evidence that the primary sequence of PrP affects PrP-res formation and susceptibility to TSE infectivity, the means by which mutations in PrP-sen lead to PrP-res and familial TSE diseases is largely unknown. Although in one type of familial TSE, both mutant and nonmutant PrP-sen can be converted to PrP-res (Chen et al., 1997), in most cases only the mutant PrP-sen is found in the protease-resistant form (Kitamoto et al., 1991; Kitamoto et al., 1993; Tagliavini et al., 1994). Thus, the currently favored hypothesis is that mutations in the PrP gene lead to a PrP molecule, which is more likely to spontaneously convert to the abnormal form than nonmutant PrP-sen. The difference between

FIG. 6. Mutations in human PrP associated with familial TSE and disease susceptibility. The structure of human PrP is shown (Zahn et al., 2000). The legend is the same as for Fig. 2A. The locations of single amino acid mutations and insertion or deletion mutants associated with human familial TSE diseases are indicated below the line (see Goldfarb et al., 1996 for review). For comparison, the location of amino acid residues associated with resistance to disease or PrP-res formation are indicated above the line.

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this type of “spontaneous” conversion and the “induced” conversion associated with exposure to infectivity is that in spontaneous conversion both PrP-sen and PrP-res are derived from within the host. 1. Properties of Mutant PrP Associated with Familial TSE Disease It is now quite clear that mutant PrP molecules associated with familial TSE diseases have properties different from wild-type PrP and that these altered properties lead to a PrP molecule with properties reminiscent of PrP-res. For example, PrP-sen with the Gerstmann-Sträussler-Schienker syndrome (GSS)-associated aspartic acid to asparagine mutation at codon 178 demonstrates an altered cell surface expression (Petersen et al., 1996), whereas the GSS-associated alanine to valine mutation at codon 117 displays an aberrant transmembrane form (Hay et al., 1987; Lopez et al., 1990; Hegde et al., 1998; Hegde et al., 1999). PrP with the CJD-associated glutamate to lysine mutation at codon 200 yields a PrP that is insoluble (Gabizon et al., 1996). Insertion of extra copies of an octapeptide repeat motif in PrP has been associated with CJD in several different families and is one of the best studied familial TSE mutations. These mutants are processed differently (Lehmann and Harris, 1995; Daude et al., 1997) and somewhat resemble PrP-res in that as the number of octapeptide repeats increases they have a greater tendency to aggregate as well as an increased resistance to proteinase K (see Harris, this volume and Lehmann and Harris, 1996; Priola and Chesebro, 1998). 2. Mutant PrP-sen vs. PrP-res Although all of these studies show that mutant PrP molecules behave differently than nonmutant PrP, there is still no clear evidence that any of the mutations shown in Fig. 6 actually induce disease by spontaneously forming PrP-res. Although PrP mutants can be slightly more resistant to proteinase K, their level of resistance is several hundredfold less than that of PrP-res isolated from the brain (Priola and Chesebro, 1998). PrP mutants containing extra copies of the octapeptide repeat region were unable to spontaneously form PrP-res and were no different from nonmutant PrP in their ability to induce PrP-res formation (Priola and Chesebro, 1998). Furthermore, although transgenic mice overexpressing one of these mutants develop neurological disease, no highly proteinase K resistant PrP-res is detectable (Chiesa et al., 1998). B. Mechanisms of Disease in Familial TSE Diseases Given the number of different PrP mutations associated with heritable TSE diseases and their varied locations in the PrP molecule (Fig. 6),

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it is possible that there is no single pathogenic mechanism common to all the familial TSE diseases. Studies using transgenic mice expressing mutant PrP molecules associated with human familial TSE diseases suggest that this may be the case. In one transgenic mouse model of GSS, overexpression of the mutant allele may be important (Hsiao et al., 1990), but in a transgenic mouse carrying a different GSS mutation, a transmembrane form of PrP may influence pathogenesis (Hegde et al., 1998; Hegde et al., 1999). Another possible mechanism that could influence pathogenesis and formation of PrP-res involves either interactions between wild-type PrP and mutant PrP (Gabizon et al., 1996; Telling et al., 1996a) or between wild-type PrP and truncated forms of PrP that have been described in both CJD and GSS (Tagliavini et al., 1991; Kitamoto et al., 1993; Tagliavini et al., 1994; Chen et al., 1995). PrP peptides have been reported to influence PrP-res formation (Kaneko et al., 1995) and a synthetic PrP peptide has been shown to induce a GSS-like disease in certain transgenic mice (Kaneko et al., 2000). Additionally, under certain conditions some PrP peptides have been shown to be neurotoxic (Forloni et al., 1993; Hope et al., 1996), suggesting that even fragments of the PrP protein may be sufficient to induce neurological damage. The caveat with all of the current transgenic mouse models of familial TSE is that none precisely replicate the human disease. Almost all of them are dependent on transgenic mouse studies that involve both overexpression and random insertion of the transgene. In fact, when a single copy of a PrP mutant associated with GSS (proline to leucine at 102) is specifically substituted into the PrP gene locus, no spontaneous neurodegenerative disease is observed (Manson et al., 1999). Rather, an increased resistance to infection with scrapie is found (Manson et al., 1999). Thus, while the implications provided by these studies concerning the mechanism of disease induction are provocative, the molecular basis for the various familial TSE diseases remains an open question. VII. CONCLUDING REMARKS PrP-res can induce its own formation (Kocisko et al., 1994), an essential prerequisite for a protein-only infectious agent. This “replication” is clearly influenced by PrP primary sequence and conformation. Supporting data are now available for how, in the absence of any nucleic acid component, these variations in PrP sequence and conformation could account for TSE species and strain characteristics. It is important to remember, however, that the link between these properties of PrP-res at the molecular level and the in vivo events in TSE disease remains

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unclear. For example, if PrP-res conformation is the basis for TSE strains, how does conformation alone determine the different neuropathologies, incubation times, and species specificities associated with different strains? How could PrP-res determine which host cells are infected or why some strains of TSE are more dependent on replication in the lymphoreticular system than others? Conversely, what contributions, other than PrP-sen, does the infected host cell make, and how do these contributions vary from cell type to cell type? As a transmissible neurodegenerative disease involving protein misfolding and amyloid formation, the TSE diseases are undoubtedly biologically unique. The question remains whether PrP-res alone can directly dictate all aspects of TSE pathogenesis or whether as yet unidentified factors or more conventional viral components are also involved.

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Miura, T., Hori-i, H., and Takeuchi, H. (1996). FEBS Lett. 396, 248–252. Moore, R.C., Hope, J., McBride, P.A., McConnell, I., Selfridge, J., Melton, D.W., and Manson, J.C. (1998). Nat. Genet. 18, 118–125. O’Conner, S.E., and Imperiali, B. (1996). Chem. Biol. 3, 803–812. O’Conner, S.E., and Imperiali, B. (1998). Chem. Biol. 5, 427–437. Palmer, M.S., Dryden, A.J., Hughes, J.T., and Collinge, J. (1991). Nature 352, 340–342. Pan, K.-M., Baldwin, M., Nguyen, J., Gasset, M., Serban, A., Groth, D., Mehlhorn, I., Huang, Z., Fletterick, R.J., Cohen, F.E., and Prusiner, S.B. (1993). Proc. Natl. Acad. Sci. U.S.A. 90, 10962–10966. Parchi, P., Castellani, R., Capellari, S., Ghetti, B., Young, K., Chen, S.G., Farlow, M., Dickson, D.W., Sima, A.A.F., Trojanowski, J.Q., Petersen, R.B., and Gambetti, P. (1996). Ann. Neurol. 39, 767–778. Parchi, P., Capellari, S., Chen, S.G., Petersen, R.B., Gambetti, P., Kopp, N., Brown, P., Kitamoto, T., Tateishi, J., Giese, A., and Kretzschmar, H. (1997). Nature 386, 232–234. Petersen, R.B., Parchi, P., Richardson, S.L., Urig, C.B., and Gambetti, P. (1996). J. Biol. Chem. 271, 12661–12668. Piccardo, P., Seiler, C., Dlouhy, S.R., Young, K., Farlow, M.R., Prelli, F., Frangione, B., Bugiani, O., Tagliavini, F., and Ghetti, B. (1996). J. Neuropathol. Exp. Neurol. 55, 1157–1163. Pocchiari, M. (1994). Mol. Aspects Med. 15, 195–291. Priola, S.A., (1999). Biomed. Pharmacother. 53, 27–33. Priola, S.A., Caughey, B., Race, R.E., and Chesebro, B. (1994). J. Virol. 68, 4873–4878. Priola, S.A., and Caughey, B. (1994). Mol. Neurobiol. 8, 113–120. Priola, S.A., and Chesebro, B. (1995). J. Virol. 69, 7754–7758. Priola, S.A., and Chesebro, B. (1998). J. Biol. Chem. 273, 11980–11985. Prusiner, S.B. (1982). Science 216, 136–144. Prusiner, S.B., Scott, M., Foster, D., Pan, K.M., Groth, D., Mirenda, C., Torchia, M., Yang, S.L., Serban, D., Carlson, G.A., Hoppe, P.C., Westaway, D., and DeArmond, S.J. (1990). Cell 63, 673–686. Prusiner, S.B. (1991). Science 252, 1515–1522. Pushkin, S.V., Kushnirov, V.V., Smirnov, V.N., and Ter-Avanesyan, M.D. (1997). Science 277, 381–383. Race, R., and Chesebro, B. (1998). Nature 392, 770. Race, R.E., Priola, S.A., Bessen, R.A., Ernst, D., Dockter, J., Rall, G.F., Mucke, L., Chesebro, B., and Oldstone, M.B.A. (1995). Neuron 15, 1183–1191. Raeber, A.J., Race, R.E., Brandner, S., Priola, S.A., Sailer, A., Bessen, R.A., Aguzzi, A., Oldstone, M.B.A., Weissmann, C., and Chesebro, B. (1997). EMBO J. 16, 6057–6065. Raeber, A.J., Sailer, A., Hegyi, I., Klein, M.A., Rulicke, T., Fischer, M., Brandner, S., Aguzzi, A., and Weissmann, C. (1999). Proc. Natl. Acad. Sci. U.S.A. 96, 3987–3992. Raymond, G.J., Hope, J., Kocisko, D.A., Priola, S.A., Raymond, L.D., Bossers, A., Ironside, J., Will, R.G., Chen, S.G., Petersen, R.B., Gambetti, P., Rubenstein, R., Smits, M.A., Lansbury, P.T., Jr., and Caughey, B. (1997). Nature 388, 285–288. Riek, R., Hornemann, S., Wider, G., Billeter, M., Glockshuber, R., and Wuthrich, K. (1996). Nature 382, 180–182. Riek, R., Hornemann, S., Wider, G., Glockshuber, R., and Wuthrich, K. (1997). FEBS Lett. 413, 282–288. Rogers, M., Yehiely, F., Scott, M., and Prusiner, S.B. (1993). Proc. Natl. Acad. Sci. U.S.A. 90, 3182–3186. Safar, J., Ceroni, M., Gajdusek, D.C., and Gibbs, C.J., Jr. (1991). J. Infect. Dis. 163, 488–494. Safar, J., Roller, P.P., Gajdusek, D.C., and Gibbs, C.J., Jr. (1993a). J. Biol. Chem. 268, 20276–20284.

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Safar, J., Roller, P.P., Gajdusek, D.C., and Gibbs, C.J., Jr. (1993b). Protein Sci. 2, 2206–2216. Safar, J., Wille, H., Itri, V., Groth, D., Serban, H., Torchia, M., Cohen, F.E., and Prusiner, S.B. (1998). Nat. Med. 4, 1157–1165. Scott, M., Foster, D., Mirenda, C., Serban, D., Coufal, F., Walchli, M., Torchia, M., Groth, D., Carlson, G., DeArmond, S.J., Westaway, D., and Prusiner, S.B. (1989). Cell 59, 847–857. Scott, M., Groth, D., Foster, D., Torchia, M., Yang, S.L., DeArmond, S.J., and Prusiner, S.B. (1993). Cell 73, 979–988. Scott, M.R., Kohler, R., Foster, D., and Prusiner, S.B. (1992). Protein Sci. 1, 986–997. Shaked, G.M., Fridlander, G., Meiner, Z., Taraboulos, A., and Gabizon, R. (1999). J. Biol. Chem. 274, 17981–17986. Smith, C.J., Drake, A.F., Banfield, B.A., Bloomberg, G.B., Palmer, M.S., Clarke, A.R., and Collinge, J. (1997). FEBS Lett. 405, 378–384. Snow, A.D., Kisilevsky, R., Willmer, J., Prusiner, S.B., and DeArmond, S.J. (1989). Acta Neuropathol. 77, 337–342. Somerville, R.A., (1999). J. Gen. Virol. 80, 1865–1872. Somerville, R.A., Millson, G.C., and Kimberlin, R.H. (1980). Intervirology 13, 126–129. Somerville, R.A., Chong, A., Mulqueen, O.U., Birkett, C.R., Wood, S.C., and Hope, J. (1997). Nature 386, 564. Somerville, R.A., and Ritchie, L.A. (1990). J. Gen. Virol. 71, 833–839. Stahl, N., Borchelt, D.R., Hsiao, K., and Prusiner, S.B. (1987). Cell 51, 229–240. Stahl, N., Borchelt, D.R., and Prusiner, S.B. (1990). Biochemistry 29, 5405–5412. Stockel, J., Safar, J., Wallace, A.C., Cohen, F.E., and Prusiner, S.B. (1998). Biochemistry 37, 7185–7193. Tagliavini, F., Prelli, F., Ghiso, J., Bugiani, O., Serban, D., Prusiner, S.B., Farlow, M.R., Ghetti, B., and Frangione, B. (1991). EMBO J. 10, 513–519. Tagliavini, F., Prelli, F., Porro, M., Rossi, G., Giaccone, G., Farlow, M.R., Dlouhy, S.R., Ghetti, B., Bugiani, O., and Frangione, B. (1994). Cell 79, 695–703. Taraboulos, A., Rogers, M., Borchelt, D.R., McKinley, M.P., Scott, M., Serban, D., and Prusiner, S.B. (1990). Proc. Natl. Acad. Sci. U.S.A. 87, 8262–8266. Taraboulos, A., Raeber, A.J., Borchelt, D.R., Serban, D., and Prusiner, S.B. (1992). Mol. Biol. Cell. 3, 851–863. Taraboulos, A., Scott, M., Semenov, A., Avraham, D., Laszlo, L., and Prusiner, S.B. (1995). J. Cell. Biol. 129, 121–132. Taylor, K.L., Cheng, N., Williams, R.W., Steven, A.C., and Wickner, R.B. (1999). Science 283, 1339–1343. Telling, G.C., Scott, M., Mastrianni, J., Gabizon, R., Torchia, M., Cohen, F.E., DeArmond, S.J., and Prusiner, S.B. (1995). Cell 83, 79–90. Telling, G.C., Haga, T., Torchia, M., Tremblay, P., DeArmond, S.J., and Prusiner, S.B. (1996a). Genes Dev. 10, 1736–1750. Telling, G.C., Parchi, P., DeArmond, S.J., Cortelli, P., Montagna, P., Gabizon, R., Mastrianni, J., Lugaresi, E., Gambetti, P., and Prusiner, S.B. (1996b). Science 274, 2079–2082. Telling, G.C., Tremblay, P., Torchia, M., DeArmond, S.J., Cohen, F.E., and Prusiner, S.B. (1997). Protein Sci. 6, 825–833. Turk, E., Teplow, D.B., Hood, L.E., and Prusiner, S.B. (1988). Eur. J. Biochem. 176, 21–30. Wells, G.A.H., Scott, A.C., Johnson, C.T., Gunning, R.F., Hancock, R.D., Jeffrey, M., Dawson, M., and Bradley, R. (1987). Vet. Rec. 121, 419–420. Westaway, D., Goodman, P.A., Mirenda, C.A., McKinley, M.P., Carlson, G.A., and Prusiner, S.B. (1987). Cell 51, 651–662.

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Wickner, R.B. (1994). Science 264, 566–569. Will, R.G., Ironside, J.W., Zeidler, M., Cousens, S.N., Estibeiro, K., Alperovitch, A., Poser, S., Pocchiari, M., Hofman, A., and Smith, P.G. (1996). Lancet 347, 921–925. Williams, E.S., and Young, S. (1980). J. Wildl. Dis. 16, 89–98. Williams, E.S., and Young, S. (1982). J. Wildl. Dis. 18, 465–471. Wong, C., Xiong, L-W., Horiuchi, M., Raymond, L., Wehrly, K., Chesebro, B., and Caughey, B. (2001). Embo J. 20, 377–386. Wopfner, F., Weidenhofer, G., Schneider, R., von Brunn, A., Gilch, S., Schwarz, T.F., Werner, T., and Schatzl, H.M. (1999). J. Mol. Biol. 289, 1163–1178. Zahn, R., Liu, A., Luhrs, T., Riek, R., von Schroetter, C., Garcia, F.L., Billeter, M., Calzolai, L., Wider, G., and Wuthrich, K. (2000). Proc. Natl. Acad. Sci. U.S.A. 97, 145–150. Zulianello, L., Kaneko, K., Scott, M., Erpel, S., Han, D., Cohen, F.E., and Prusiner, S.B. (2000). J. Virol. 74, 4351–4360.

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MASS SPECTROMETRIC ANALYSIS OF PRION PROTEINS BY MICHAEL A. BALDWIN Mass Spectometry Facility, University of California, San Francisco, California 94143

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

VII. VIII. IX. X. XI.

Modern Mass Spectrometric Techniques for Protein Characterization . . . . Identification and Preliminary Analysis of PrP . . . . . . . . . . . . . . . . . . . . . . . . Confirmation of the PrPSc Amino Acid Sequence. . . . . . . . . . . . . . . . . . . . . . Non-PrP Peptides in Prion Preparations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . N -linked Oligosaccharides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Analysis of the GPI Anchor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. ESIMS of the C-terminal Peptide-GPI. . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Branching Patterns in the GPI Glycan by MS/MS . . . . . . . . . . . . . . . . . . C. Unanswered Questions Concerning the GPI . . . . . . . . . . . . . . . . . . . . . . Analysis of Intact PrP by MALDIMS and ESIMS . . . . . . . . . . . . . . . . . . . . . . . Processing of Chicken PrP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Recombinant PrP and Synthetic Peptides. . . . . . . . . . . . . . . . . . . . . . . . . . . . A. PrP Modifications Due to Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Copper Binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Accessory Molecules in Scrapie Prions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

29 30 31 37 37 39 41 42 43 45 49 49 50 50 51 51 52

1. MODERN MASS SPECTROMETRIC TECHNIQUES FOR PROTEIN CHARACTERIZATION As a result of developments in the methods for desorption and ionization of polar and labile materials from condensed phase, mass spectrometry has become a powerful tool for biochemical analysis. In the 1980s the new methods of fast atom bombardment mass spectrometry (FABMS) (1) and liquid secondary ionization mass spectrometry (LSIMS) (2) allowed the analysis and sequencing of peptides without derivatization, and the detection and identification of posttranslational modifications (3). These methods enabled the routine, rapid, and accurate mass analysis of peptides isolated from proteolytic or chemical digestions, complementing longer established chemical techniques such as amino acid analysis and Edman N-terminal sequencing. More recently these methods were supplanted by electrospray ionization (ESI) (4, 5) and matrix assisted laser desorption/ionization (MALDI) (6–9). In addition to being applicable to peptides, these latter techniques have proved to be capable of ionizing large intact proteins and providing highly accurate molecular masses, in favorable cases with 29 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved. 0065-3233/01 $35.00

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parts-per-million accuracy and with subpicomole sensitivity. They are also applicable to the analysis of nucleic acids including DNA and RNA, oligosaccharides, and lipids (10). Much of the success of these so-called “soft ionization” methods results from the low levels of vibronic excitation imparted to the molecules during the ionization process. Thus molecular ions are less likely to break down and they are mostly preserved intact, even for labile and potentially unstable compounds. Structural information can be obtained by collision-induced dissociation (CID) of these preponderant molecular ions in a tandem mass spectrometer, which can lead directly to the sequence of amino acids in a peptide or the sequence and branching of sugar units in an oligosaccharide. This technique is also referred to as mass spectrometry/mass spectrometry (MS/MS). Today mass spectrometry is capable of determining the amino acid sequence and posttranslational modifications of virtually any protein that can be purified in sufficient quantity. II. IDENTIFICATION AND PRELIMINARY ANALYSIS OF PrP After PrP was first cloned in the mid-1980s (11), it was soon established that PrPSc and PrPC are both encoded by the same cellular gene and that PrPSc is derived from PrPC in a posttranslational event (12, 13). Despite dramatic differences in the physical properties of PrPC and PrPSc, biochemical analysis failed to identify any chemical differences. Both forms were shown to contain a single disulfide bond (14). They are modified with N-linked oligosaccharides that cause intact PrP to give three separate bands by SDS-PAGE at 33–35 kDa and that can be removed with peptide N-glycosidase F (PNGase F) (15, 16). Both forms bind to cellular membranes via a glycosyl phosphatidylinositol (GPI) group, the lipids of which can be removed by phosphatidylinositol phospholipase C (PIPLC) (17). A series of articles published from the University of California, San Francisco, between 1990 and 1993 described the use of mass spectrometry to address questions that might explain the pathological properties of PrPSc. Do processes such as RNA-editing cause changes in the amino acid sequence? Are there differences in posttranslational modifications of PrPC and PrPSc? Do any identified modifications possess unusual features that could explain the pathogenic nature of PrPSc?

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Is the protein homogeneous or could a subpopulation be responsible for the unusual properties? It is noteworthy that although mass spectrometry is now widely used for the identification and characterization of proteins, it is quite unusual to carry out a complete analysis in which every amino acid of a protein is probed. Measuring the molecular weights of a subset of peptides from a proteolytic digest or determining a region of amino acid sequence is normally sufficient to identify a protein present in a database. For previously unknown proteins or those for which no corresponding genetic sequence is available, some amino acid sequence will normally allow cloning and expression of recombinant products. If such a protein has the “correct molecular weight,” it is generally assumed to be the target protein, particularly if it has the same activity, although a protein expressed in bacteria is likely to lack the posttranslational modifications of the natural form. In the case of PrP, such an approach was not adequate to answer the questions posed previously. At the time there were well-established methods for purifying proteolytically truncated PrPSc from the brains of hamsters artificially infected with scrapie. PrPSc accumulates during disease and its insolubility and unusual resistance to proteolysis allow a truncated core (PrP 27–30) to be isolated after treatment with proteinase-K (18–20). This retains infectivity, despite the loss of up to 67 amino acids from the N-terminus. Much of the analysis was carried out using this form, although the full-length protein was also isolated in lower yield for analysis of the N-terminal region. Both forms required the presence of detergent for solubilization, and they were rendered more soluble by chaotropic agents such as 6 M guanidine hydrochloride. Although guanidine denatured the protein and destroyed the infectivity, it did not diminish the effectiveness of mass spectrometric determination of the primary structure. Before analysis the disulfide bond was reduced with dithiothreitol and carboxymethylated with iodoacetic acid. The GPI lipids were incompatible with reversed phase high-performance liquid chromatography (HPLC), consequently they were removed by digestion with PIPLC. PrPC was more difficult to isolate and at that time was only available in very small quantities (21). III. CONFIRMATION OF THE PrPSC AMINO ACID SEQUENCE The Syrian hamster PrP gene sequence was already known from partial purification of PrP 27–30, Edman N-terminal sequencing, cloning of a cDNA, and subsequent similar studies on full length PrPSc and

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PrPC. These methods revealed a 254-amino acid sequence from which an N-terminal 22-amino acid hydrophobic signal peptide is removed in the development of the mature proteins, which start at Lys23. The major start site for PrP 27–30 was known from Edman sequencing to be Gly90. The C-terminus was also known to be modified by the removal of another hydrophobic peptide of unknown length and the attachment of the GPI anchor. SDS-PAGE mobilities had shown that the majority of protein molecules carry oligosaccharides at both the consensus sites, Asn181 and Asn197. To confirm that the sequence of PrP was identical to that predicted from the gene sequence, it was necessary to cleave it selectively at specific residues to give a number of smaller peptide fragments. Hopefully these could be separated by HPLC and analyzed either off-line by collecting fractions or on-line by directly coupled HPLC-MS. The strategy adopted was to select an enzyme that would produce a relatively small number of peptides, the smaller of which would be analyzed directly and the larger would be digested with a second enzyme for further analysis. Originally it was intended to analyze all of the peptides by LSIMS, which in routine use has optimal sensitivity below 2500 Da but during the course of the work the further development of ESIMS allowed the direct analysis of much larger peptides. Endoproteinase Lys-C cleaving at the C-terminal side of lysine was selected as the first enzyme, for which PrP 27–30 should give nine fragments and full length PrP should give twelve fragments ranging from single lysines to a 75 amino acid glycopeptide His111-Lys185. In practice not all sites were cleaved with equal efficiency, (e.g., Lys-Pro bonds, of which there are three in PrP, are relatively stable and digestion was incomplete). Lys-C retains its activity in 0.1% sodium dodecyl sulfate (SDS) which is required to keep the protein in solution, even after denaturation and after removal of the GPI lipids with the enzyme PIPLC. SDS was subsequently removed by precipitation with guanidine as it interferes with most chromatographic and mass spectrometric methods. An HPLC chromatogram of a Lys-C digest of PrP 27–30 is illustrated in Fig. 1. LSIMS involves the ablation of cationized species such as protonated molecular ions from solution in a weakly acidic viscous liquid matrix of low volatility by bombardment with high energy cesium ions. This is very similar to FABMS, which uses argon or xenon atoms rather than cesium ions. A common matrix for both techniques is 1:1 glycerol/thioglycerol with 0.1% trifluoroacetic acid (TFA). In this study the ions were mass analyzed in a double-focusing sector mass spectrometer operating at ~3000 resolving power; thus peaks at adjacent mass numbers were clearly resolved, allowing mass measurement to ±0.2 Da. LSIMS was ideal for studying the midsize peptides of about 5 to 20

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FIG. 1. Reversed phase HPLC chromatogram of an endoproteinase Lys C digest of PrP 27–30, showing absorbance at 214 and 280 nm versus time (min). The gradient line shows acetonitrile content. The peptide compositions shown for each peak were derived from the experiments described in Section III. Peptides were identified containing all residues between 74 and 231, with the exception of an anticipated tetrapeptide, residues 107–110. (Reproduced with permission from Stahl, N. et al. in Prusiner, S.B., Collinge, J., Powell, J., and Anderton, B. [1992], Prion Diseases of Humans and Animals, pp. 361–379. Copyright Elsevier Science).

residues, seven of the nine predicted fragments from PrP 27–30 falling in this category. The 75 residue peptide was anticipated to be too large for this methodology and the tetrapeptide Thr-Asn-Met-Lys might be too small and hydrophilic for efficient detection. The primary information provided by mass spectrometry is molecular mass, measurements being made on each peak eluting from the HPLC. From the predicted protein sequence a computer program generated all possible cleavage products for any given enzyme, matching the measured and calculated masses. Any HPLC fraction giving a positive match could be further analyzed by amino acid analysis (AAA), MS/MS and/or Edman N-terminal sequencing. In this way the majority of the PrP peptides were identified and confirmed to have the sequences anticipated from the cDNA sequence (22). Some modifica-

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MICHAEL A. BALDWIN

tions were observed. An early eluting fraction gave a strong peak at m/z 1016.3, in agreement with the calculated mass of the protonated peptide Gln186-Lys194. A slightly later fraction with the same composition by AAA gave a peak at m/z 999.3. The latter peptide gave no signal by Edman analysis, indicating a blocked N-terminus as was confirmed by MS/MS, which showed the first residue had been converted to pyroglutamic acid (23). The peptide corresponding to residues 195–204, Gly-Glu-Asn-PheThr-Glu-Thr-Asp-Ile-Lys was anticipated to be glycosylated at Asn197. A fraction isolated as a weak HPLC peak gave a signal at m/z 1153.5, corresponding to the peptide without oligosaccharide, which was confirmed by both MS/MS and Edman. A larger and slightly earlier eluting HPLC peak had the same amino acid analysis but gave no signal by LSIMS. Edman analysis confirmed the sequence except that there was no signal at the third cycle, presumably due to glycosylation of the asparagine. Treating the peptide with PNGase F removed the N-linked oligosaccharide and converted Asn197 to aspartic acid. This was observable by LSIMS as a 1 Da increase in the peptide m/z, in this instance to m/z 1154.5. MS/MS showed that the first two residues were unchanged, the so-called b2 ion for Gly-Glu being at m/z 187. The b3 ion moved from m/z 301 for Gly-Glu-Asn for the naturally unglycosylated peptide to m/z 302 for Gly-Glu-Asp of the PNGase F-treated glycopeptide, as shown in Fig. 2. A minor component (~15%) was identified at m/z 932.3 as a C-terminal peptide Glu221-Gly228 having no GPI anchor. This was not an anticipated Lys-C cleavage site and was attributed to a small fraction of PrP that was devoid of the GPI (24). A further HPLC fraction containing the major C-terminal peptide carrying the GPI group was tentatively identified by AAA, although this gave no signal by LSIMS. Potential chemical and enzymatic methods to simplify this complex structure are summarized in Fig. 3. The GPI was removed by overnight treatment with 50% HF at 4°C, which hydrolyzes any phosphodiester linkages or phosphate groups (25). LSIMS then gave the mass of the protonated peptide as 1354.5 Da, corresponding to Glu221-Ser231, the C-terminal serine carrying the residual ethanolamine remaining after hydrolysis of the GPI. Thus, consistent with earlier predictions (25), the previously unknown site of attachment of the GPI was identified as Ser231 (24). Experiments with synthetic peptides established that the hydrophilic tetrapeptide Thr-Asn-Met-Lys was not retained on the C18 HPLC column. The HPLC column flow-through from the Lys-C digest was derivatized to add a cholate group to the N-terminus. The increased hydrophobicity allowed the derivatized peptide to be repurified by HPLC and analyzed by

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FIG. 2. A portion of the tandem mass spectrum of the glycopeptide, residues 195–204, containing the second glycosylation site, after treatment with PNGase F. The mass difference of 115 Da between the b2 and b3 ions is attributable to aspartic acid as the third residue (codon 197) rather than asparagine. (Reproduced with permission from Baldwin, MA. et al. [1993]. Trends in Analytical Chemistry 12, 239–248. Copyright Elsevier Science.)

LSIMS. The positive charge of the cholate group also enhanced the LSIMS signal, which was observed at the predicted m/z 652.3 (22). In addition to experiencing difficulties with the C-terminal peptideGPI, there were two peptides that were too large to be directly amenable to analysis by LSIMS, the 75 residue peptide of PrP 27–30 and a large Nterminal portion of full-length PrPSc. However, about this time ESIMS and MALDIMS became available for the analysis of such peptides. ESIMS was carried out by injecting the analyte into a flowing liquid matrix of 1:1 acetonitrile/water containing 1% acetic acid, which was sprayed from a positively charged needle into a chamber at atmospheric pressure. The resulting charged droplets evaporated under the influence of a stream of nitrogen. Analyte molecules were multiply protonated by the acid medium and drawn through an aperture into the vacuum of the mass spectrometer. Because mass spectrometers separate according to mass/number of charges (m/z), it was possible to monitor highly charged ions with molecular weights greater than the nominal mass range of the mass spectrometer. Furthermore, isolated molecular ions were obtained in the gas phase without the requirement for the analyte to be volatile. Lys C digestion of the Lys-Pro bonds at Lys27, Lys101, and Lys104 was incomplete; and the N-terminal portion from full length PrPSc was man-

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FIG. 3. Chemical and enzymatic treatment to reduce the size and complexity of the GPI anchor for mass spectrometric analysis. PIPLC removed the acylalkylglycerol lipids, then endoproteinase Lys C cut the amino acid chain after Lys220, giving the C-terminal peptide attached to the phosphorylated glycan. Incubation with 50% aqueous HF was used to hydrolyze the phosphodiester bonds and to release the glycan and the peptide, which were separated by RP-HPLC and analyzed independently.

ifested as a series of closely related peptides poorly resolved by HPLC. Four peptides were successfully identified by ESIMS, with a mean difference between the predicted and measured masses of 1.2 Da or 0.015% (22). The largest was Arg25-Lys106 of molecular mass 8,294.8 Da, observed as a series of ions with between seven and thirteen charges in the range m/z 600 to 1200. Further digestion by trypsin cleaved at Arg37 and Arg47, allowing smaller peptides to be isolated and analyzed. Arg25 was found to be resistant to trypsin as it is followed by proline. Earlier Edman sequencing of the intact protein had suggested possible modification to Arg25 and Arg37, but this was not confirmed in the present study. It was subsequently suggested that the modification might be to citrulline (J. Hope, personal communication), the mass of which is only 1 Da higher than arginine. Because the purification of full-length PrPSc was less effective than that of PrP 27–30, the quality of the original data for the full length peptide made it difficult to distinguish unambiguously between the possibilities of arginine or citrulline at these two positions. However, trypsin would not digest at citrulline; therefore the observation of the expected shorter peptides in the trypsin digest con-

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firmed that at least a fraction of the molecules contained arginine rather than citrulline at residue 37. The tryptic peptide of residues 23–37, presumably containing Arg25, gave the expected mass for the presence of arginine rather than citrulline, although a substoichiometric amount of citrulline could be present, as the M+1 ion was more intense than would be expected based on stable isotope ratios. IV. NON-PrP PEPTIDES IN PRION PREPARATIONS In the purifcation of PrPSc from brain tissue, it was very difficult to eliminate all other proteins. Every PrPSc preparation gave peptide signals that could not be attributed to the PrP sequence. The contaminants were often at low level and frequently coincided in HPLC fractions with PrP peptides. The HPLC fraction containing the N-terminal peptide of PrP 27–30 Gly90-Lys101 (m/z 1283.5) from several different preparations was always accompanied by ions at m/z 772.3 and 915.4. Edman analysis confirmed the presence of a mixture of peptides but could not distinguish signals from the different components and gave only limited sequence data. By contrast MS/MS was used to study each individual component in the mixture, characterizing these particular peptides as Asp-Gly-Pro-Arg-Leu-Ser-Lys and Arg-Glu-Ile-Val-AspArg-Lys. The first of these showed some homology with the leucine zipper region of a Drosophila protein, and the second showed some homology to mouse actin but the hamster sequence had not been reported. It is unlikely that these or other unidentified peptides play any role in the development of scrapie. V. N-LINKED OLIGOSACCHARIDES Preliminary studies by Endo et al. identified some of the basic structures of the N-linked oligosaccharides (15). The ESIMS analysis of the peptide His111-Lys185 confirmed that it is modified by the addition of heterogeneous groups that increase the molecular mass by 1800 to 3000 Da. Figure 4A shows the ESIMS spectrum with a repeating cluster of ions carrying 7–10 charges. This was expected as this peptide includes the first glycosylation site at Asn181. This spectrum was transformed to convert the multiply charged ion profiles into molecular masses (Fig. 4B). After treatment with PNGase F a single component was obtained (not shown), the molecular mass of which was 8607.8 Da as measured by ESIMS, close to the predicted value of 8608.6 for the deglycosylated peptide with con-

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FIG. 4. ESIMS spectrum of the glycopeptide His111-Lys185. (A) Raw data showing clusters of multiply charged species with 7–10 protons. (B) Transformed spectrum showing molecular masses rather than m/z, with three predominant species of 10256, 10564, and 10873 Da. (Reproduced with permission from Baldwin, M.A. et al. [1993]. Trends in Analytical Chemistry 12, 239–248. Copyright Elsevier Science.)

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version of asparagine to aspartic acid (22). The differences between the masses before and after treatment with PNGase F, as measured both by ESIMS and MALDIMS, confirmed the identity of the major oligosaccharides that had been partly characterized in the previous study (15). Thus the most abundant glycosylated species of mass 10564.5 Da has an oligosaccharide of measured mass 1956.3 Da, within 0.3 Da of that calculated for the structure of composition Hex4.HexNAc5.Fuc2 previously identified as the major neutral sugar. A more recent comprehensive mass spectrometric study of the N-linked sugars of murine PrPSc involved separation of the two glycosylation sites by tryptic digestion (26). Glycopeptide mixtures from sequential exoglycosidase digestions with combinations of neuraminidase, alpha-mannosidase and β galactosidase, were characterized by HPLC-ESIMS and MS/MS. This identified approximately 60 different sugar structures on PrPSc and demonstrated the presence of larger and more complex sugars at Asn196 than at Asn180 (mouse numbering), as shown in Fig. 5. A separate study that compared the sugars of hamster PrPC and PrPSc involved a completely different methodology in which the sugars were released from the protein, N-acetylated, fluorescently labeled by reductive aminolysis, separated by chromatography and analyzed by MALDIMS (27). It was found that essentially the same array of sugars was present in both isoforms, even though the structures containing bisecting N-acetylglucosamine residues were diminished in abundance in hamster PrPSc relative to PrPC. This significant result implies that PrPSc is not derived from any special subset of specifically glycosylated PrPC, although the reduced levels of GlcNAc imply a decrease in the activity of N-acetyaminotransferase III in scrapieinfected cells. Coincidently, these two studies also showed the sugars of mouse and hamster PrPSc to be essentially the same. VI. ANALYSIS OF THE GPI ANCHOR PrP was first identified as a GPI-anchored protein by Stahl et al. (17). All GPI anchors that have been fully characterized conform to a common pattern with a sugar glycan core containing the sequence Man3.HexN.Inos phosphate, the nonreducing terminus of which is attached through phosphoethanolamine to the C-terminus of the protein. The inositol phosphate has a lipophilic tail containing acylalkylglycerol. There is variability in the attachment of further sugar units and mammalian anchors have a further phosphoethanolamine. The previously utilized analytical methodology for GPI anchors using radiolabeling by hydrazinolysis, exoglycosidase digestion and gel permeation

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chromatography was not sufficiently sensitive for the small scale analysis required for PrP; thus several new approaches were implemented during this study (28–30). A. ESIMS of the C-terminal Peptide-GPI The C-terminal peptide could not be analyzed by LSIMS whilst still attached to the GPI, whereas ESIMS of an aliquot of the unhydrolysed fraction revealed several molecular species (Fig. 6). Certain components of the GPI could be predicted by analogy with other known structures (i.e., the glycan core would be linked to the protein through phosphoethanolamine, there would probably be a second phosphoethanolamine group, and the core would contain at least three mannose residues linked to glucosamine and phosphoinositol). Acylalkylglycerol had been removed by the action of PIPLC. The mass spectrometric analysis of oligosaccharides differs from that of peptides and proteins for which the basic building blocks are the 20 naturally occurring amino acids, each having a unique mass. The basic units of oligosaccharides are mostly isomers of only four or five unique compositions; thus it is relatively easy to assign the composition of an oligosaccharide on the basis of the number of hexose units (mass 162), hexosamines (mass 161), Nacetylhexosamines (mass 203), sialic acids (mass 291), etc. The various molecular species revealed by ESIMS differing in mass by combinations of 162 and 291 Da were assigned carbohydrate compositions. The smallest was calculated to contain the peptide, two phosphoethanolamines, three hexoses, one hexosamine, one N-acetyl hexosamine and phosphoinositol (i.e., the basic mammalian GPI with an additional Nacetylhexosamine). Larger forms contained a further one or two hexose units and unexpectedly, sialic acid (N-acetylneuraminic acid), a sugar not previously reported as a component of a GPI (30). Thus mass spectrometry showed the GPI was heterogeneous with at least five separate species and its composition was consistent with known GPI structures but with sialic acid as a novel component. The heterogeneity was confirmed by various separative techniques including capillary electrophoresis, Dionex high performance anion exchange chromatography (HPAEC), and reverse phase HPLC at pH 7. Cleavage FIG. 5. Oligosaccharide structures for mouse PrPSc at Asn180 and 196 (equivalent to 181 and 197 in hamster and human), obtained by Stimson et al. (26), grouped as (I) biantennary, (II) triantennary, and (III) tetraantennary. Squares, N-acetylglucosamine; open circles, mannose; closed circles, galactose; triangles, fucose; ovals, sialic acid. (Reproduced with permission from Stimson, E. et al. [1999]. Biochemistry 38, 4885–4895. Copyright American Chemical Society.)

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FIG. 6. Transformed ESIMS spectrum of the heterogeneous GPI species attached to the C-terminal peptide Glu221-Ser231. A, (Peptide 221–231).Ea.P.Hex3.(HexNAc) (P.Ea).HexN.Ino.P, [Ea = ethanolamine; Hex, hexose; HexN, hexosamine; HexNAc, Nacetylhexosamine; I, inositol; P, phosphate; Sia, sialic acid.]

of sialic acid by neuraminidase was also demonstrated. ESIMS showed that only species with at least four hexose units were sensitive to Jack bean α mannosidase and then only one residue was removed, consistent with the third mannose away from the glucosamine being the site of attachment to the protein. Mass spectrometry was not able to identify the particular isomeric form of each sugar unit (e.g., hexoses can be glucose, mannose or galactose, differing in the configuration of the ring hydroxyl groups). However, the sugars were identified by acid hydrolysis of the GPI and separation of the various monosaccharides by HPAEC. Comparison with the monosaccharide standards confirmed 3 or 4 mannose units, the glucosamine and sialic acid. N-acetylgalactosamine and galactose were also identified (30). B. Branching Patterns in the GPI Glycan by MS/MS Oligosaccharides differ from peptides and proteins in that they are frequently branched. Branching patterns are usually determined by digestion with glycosidases with specific activities followed by chromatographic analysis of the products, comparing retention times with those of standards. However, standards may not be available for studies on novel structures, and this approach requires relatively large amounts of material. Nuclear magnetic resonance analysis is even less sensitive. By contrast mass spectrometry is extremely sensitive and it was anticipated that the fragmentation induced by CID might reveal the branching pat-

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terns. MS/MS also has the ability to study individual selected species in unseparated mixtures such as the heterogeneous GPI glycans. High-resolution MS/MS with high energy CID was carried out on the glycans released by 50% HF and permethylated to enhance surface activity and volatility and to give a positively charged quaternary ammonium cation as the glucosamine became triply methylated, greatly enhancing the sensitivity for mass spectrometric detection. After methylation the masses of the protonated molecular ions for the most abundant species were measured as 1312.5, 1557.6, and 1761.6 Da, corresponding to the permethylated glycans lacking sialic acid. The MS/MS spectra shown in Fig. 7 represent fragmentation of these three species, separated in the first mass analyzer of the tandem mass spectrometer and caused to undergo collisions with neutral gas atoms, the fragment ions being separated in the second mass analyzer. The key to the interpretation of the tandem spectra was a series of ring cleavages across the individual hexose residues (so-called X-ions) that terminated in an ion of m/z 510 for the glucosamine.inositol moiety. Ions representing the loss of successive sugars revealed which sugar was substituted (i.e., which was the branch site). Complementary information came from the oxonium ions (called B ions by analogy with the fragmentation of peptide ions) from charge-remote fragmentations giving single or multiple sugar units from the nonreducing end of each chain (28). This is shown in Fig. 8 for the largest GPI species having sialic acid, or N-acetylneuraminic acid (NANA). The oxonium ions at m/z 376.1, 580.2 and 825.3 confirmed the presence of a linear sequence of ions corresponding to NANA.Hex.HexNAc. Thus tandem mass spectrometry together with chromatographic and glycosidase data allowed the identification of the six species shown in Fig. 9 (30). The branching was clearly revealed by cleavages along each branch of a biantennary structure. One branch represented the normal GPI glycan core with a chain of either three or four hexose units attached to the GlcN.lnos. The branch point was at the hexose immediately adjacent to the glucosamine with a chain of one, two, or three sugars in the sequence Sia.Glc.GlcNAc. The calculated molecular masses listed in Figure 9, which include the mass of the C-terminal peptide, can be related to the experimentally determined values in the ESI spectrum in Fig. 7. C. Unanswered Questions Concerning the GPI A further difference between oligosaccharides and proteins arises from the presence of alternative sites for linking individual sugars together. The glycosidic bond from carbon-1 of the nonreducing residue in a disaccha-

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FIG. 7. Partial tandem mass spectra for the region above m/z 500 of permethylated GPI anchor glycans devoid of sialic acid, showing the fragmentation of m/z 1312, 1557 and 1761. (Reproduced with permission from Baldwin, M.A. et al. [1990]. Analytical Biochemistry 191, 174–182. Copyright Academic Press.)

ride can be linked to positions 2, 3, 4, or 6 of the reducing residue. Furthermore, the glycosidic bond at carbon-1 has two possible configurations giving either α or β anomers. Each of these differences may be crucial in terms of biochemical action, but mass spectrometry cannot be applied easily to their analysis. Permethylation and hydrolysis followed by peracetylation and reduction with sodium borodeuteride gives partially methylated alditol acetates specifically labeled at one end, which can be analyzed by GC-MS to reveal the linkage positions. Unfortunately this strategy is difficult to prosecute with less than 5 to 10 nmole of homogeneous starting material. There are glycosidases that are specific for both anomericity and linkage position that are valuable in dealing with a structure likely to conform to a previously identified pattern. In this way it was shown that the third mannose in the chain, the site of attachment of the GPI to the protein, becomes susceptible to hydrolysis by alpha-mannosi-

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FIG. 8. Tandem mass spectrum of a permethylated GPI glycan of m/z 2123, containing sialic acid (NANA). (Reproduced with permission from Baldwin, M.A. et al. in Prusiner, S.B., Collinge, J., Powell, J., and Anderton, B. [1992]. Prion Diseases of Humans and Animals, pp. 380–397. Copyright Elsevier Science).

dase only after HF treatment. Interestingly the middle mannose in the chain of three at the core of the GPI does not show the same sensitivity (30), even though previous studies on GPIs from other proteins showed α1–6 linked mannose at this point (31). VII. ANALYSIS OF INTACT PrP BY MALDIMS AND ESIMS Enzyme digestion and peptide mapping are powerful methods for identifying the sites and the nature of any posttranslational modifications. However, measuring the intact molecular mass of the protein is the most effective way of ensuring that nothing was overlooked during the mapping experiments. ESIMS and MALDIMS are both capable of ionizing and accurately determining the mass of intact proteins. ESIMS generally offers higher resolution but for heterogeneous samples such as multiply glycosylated proteins, the presence of overlapping series of highly charged ions complicates the spectral analysis. MALDIMS is unable to resolve all the individual molecular species for a protein such as PrP, but it can provide the centroid or peak top mass of the unresolved peak profile. Samples for MALDIMS are dissolved in a solution containing a 104 to 105 molar excess of a UV-absorbing matrix, normally an aromatic acid

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FIG. 9. Structures of the various GPI anchors derived from experiments described in Section VI. The calculated masses include the C-terminal peptide, and can be directly correlated with the masses shown in Figure 5. Man, mannose; Gal, galactose; GalNAc, Nacetyl galactosamine; GlcNH2, glucosamine; Ino, inositol; Sia, sialic acid. (Reproduced with permission from Stahl, N. et al. [1999]. Biochemistry 31, 5043–5053. Copyright American Chemical Society.)

with additional conjugated double bonds such as 4-hydroxy-α-cyanocinnamic acid (4-HCCA). Without detergents or denaturants that suppress the ionization process, membrane proteins such as PrP are generally insoluble in solvents that are suitable for the matrix compound. PrPSc

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was treated with PNGase F to remove the N-linked sugars and reduce the heterogeneity, then dissolved in hexafluoroisopropanol and mixed 1:1 with a solution of 5 mg/mL 4-HCCA in 2:5:2 chloroform/ methanol/0.1% TFA. One µL of this mixture containing approximately 1 pmol of protein was deposited on the MALDI target and the solvent was evaporated. In the B. T. Chait’s laboratory at Rockefeller University, this was irradiated and ionized with 354 nm radiation from a Nd-YAG laser, the ablated ions being mass analyzed by time of flight mass spectrometry. The mass spectrum showed broad peaks corresponding to singly, doubly, and triply charged ions, and gave an average mass of ~25,350 Da (32). Taking account of the relative abundance of the different GPIs, the molecular masses for amino acids Lys23-Ser231 combined with the various GPI species (Fig. 9) and the likely GPI lipids, a weighted mean of 25,329 Da was calculated. Thus there was relatively good agreement between the calculated and measured numbers, suggesting that no major modification had been destroyed or overlooked. However, the resolving power of MALDIMS was not sufficient to separate the various forms present owing to the heterogeneity of the GPI anchor, hence the difficulty of measuring the molecular weight with high precision. ESIMS gives higher resolution, but it relies on the protein being soluble; PrPC is more soluble than PrPSc, and Fig. 9 compares more recent spectra of PrPC obtained by MALDIMS and ESIMS. Despite the removal of the sugars with PNGase F to reduce the overall heterogeneity, the heterogeneous GPI glycan remained attached. The protein was also treated with PIPLC to remove the lipids, thereby enhancing solubility in the ESIMS buffer of 1:1 acetonitrile water with 1% acetic acid. The upper spectrum shows the MALDIMS data obtained with a commercial mass spectrometer at UCSF for hamster brain PrPC mixed with myoglobin as an internal mass calibrant. The PrP peaks are broader than those of myoglobin and show unresolved structure. Using peak tops to define the mass of each ion, PrPC appeared as a singly charged ion at 24,466.1, doubly charged at 12,236.9 and triply charged at 8,161.6. The mean molecular mass calculated from these three species was 24,472.9 Da. Note that it is generally more accurate to use centroids rather than peak tops, but not in the case of unresolved peaks arising from ions of different compositions. Because of much higher charge states, a spectrum of the same sample from ESIMS in the lower panel appears to be completely different. Peaks are observed for the attachment of between 20 and 32 protons, each charge state showing at least two major species. The raw data on the left was deconvoluted to the molecular weight pattern shown in the right hand panel. Here the different glycoforms of the GPI are well resolved and give a molecular weight for the major species of 24,474.0. The calcu-

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lated value for the PrP sequence plus the major GPI glycoform is 24,474.5 Da, in excellent agreement with the ESIMS result (0.002%) and in surprisingly good agreement with MALDIMS, considering the poorly resolved nature of the MALDI spectrum. VIII. PROCESSING OF CHICKEN PrP Chicken PrP shows moderate homolgy to mammalian prion proteins. Harris et al. (33) used sequence-specific antibodies to immunoprecipitate and immunoblot chicken PrP derived from stably transfected cultures of neuroblastoma cells, as well as from chicken brain and cerebrospinal fluid. They used MALDIMS to characterize the protein fragments indicative of natural processing sites. The majority of chicken PrP protein molecules present in neuroblastoma cells and on isolated brain membranes are attached to the cell surface by the GPI anchor. Most of these surface-anchored molecules were found to be truncated at their N-termini, distal to the proline/glycine-rich repeats. Corresponding N-terminal fragments found in the medium were identified by mass spectrometry, such as a peptide extending from Lys25 to Phe116. Such cleavages were localized within a region of 24 amino acids that is identical in chPrP and mammalian PrP, and represents a major processing event that could have physiological as well as pathological significance. An alternative cleavage site within the GPI anchor caused the release of a fraction of the membrane-bound protein into the medium. IX. RECOMBINANT PrP AND SYNTHETIC PEPTIDES There have been numerous reports of the use of mass spectrometry to analyze recombinant PrP and synthetic peptides corresponding to regions of the PrP sequence. Most of these reports have not been reviewed here, as the methodolgy is well established and in general has been used purely to confirm the identity of the species.

FIG. 10. Mass spectra of PrPSc from MALDIMS, and from ESIMS. For MALDIMS the mass scale was calibrated with myoglobin ions as an internal standard. For ESIMS the mass scale was calibrated externally using the doubly charged and singly charged ions of gramicidin S. The right hand panel of the ESIMS spectrum shows the deconvoluted spectrum, giving measured molecular weights for the major species, which differ by one hexose. Deviations from the calculated values are shown in parentheses.

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A. PrP Modifications Caused by Aging Prion diseases are associated with aging, and the incidence of certain protein modifications is known to increase with age. One such modification is the conversion of asparagine to aspartic acid and isoaspartic acid. It is possible that the conversion of PrPC to PrPSc might be enhanced by the presence of such modified amino acids. After aging recombinant mouse PrP at 37°C, 0.8 mole of isoaspartyl residue per mole of protein was detected by a protein-l-isoaspartyl methyltransferase assay, with a half-life for conversion of 30 days–1. Digestion of the modified PrP with endoproteinase Lys C, followed by mass spectrometry and Edman degradation, identified Asn108 in the amino-terminal flexible region to be partially converted to aspartic acid and isoaspartic acid. A second modification was the partial isomerization of Asp226 (34). However, despite an independent search for such modifications in PrPSc, they have yet to be identified in significant amounts (35), although it remains possible that such modifications in substoichiometric numbers of PrP molecules could help to initiate the PrPSc formation or stabilize PrPSc polymers in vivo. B. Copper Binding PrP-knockout mice apparently develop normally and are noteworthy only for their resistance to infection by prions; thus the normal function of PrPC is unclear. Nevertheless, there is an increasing body of evidence that it is involved in either transport or storage of copper. One of the first pieces of evidence was the demonstration by MALDIMS and ESIMS that synthetic peptides containing multiple copies of the PrP octarepeat ProHisGlyGlyGlyTrpGlyGln selectively bind copper(II) ions, but not other divalent cations (36, 37). Mass spectrometry, particularly ESIMS, is increasingly accepted as a valid means for studying noncovalent associations, including those between proteins and metal ions (38, 39). ESIMS has been used to demonstrate that the stoichiometries of the complexes are pH dependent: a peptide containing four octarepeats chelates two Cu2+ ions at pH 6 but four at pH 7.4. At the higher pH, the binding of multiple Cu2+ ions occurs with a high degree of cooperativity, particularly for peptides extended beyond the octarepeats to incorporate His96. Dissociation constants for each Cu2+ ion binding to the octarepeat peptides were shown to be in the nanomolar to micromolar range (40). PrPC is known to concentrate in cell surface lipid-rich caveolae, which can be endocytosed as endosomes and secondary lysosomes at reduced pH. Thus PrP could function as a Cu2+ transporter, binding Cu2+ ions from the extracellular medium under

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physiological conditions and then releasing some or all of this metal within the cell on exposure to acidic pH. X. ACCESSORY MOLECULES IN SCRAPIE PRIONS Molecular biological and transgenic mouse experiments have demonstrated virtually beyond doubt that an isoform of PrP is the pathogenic agent in the TSEs. However, there have long been questions as to whether an accessory molecule is another essential component of infectious prions. One justification for this question is the failure to either regenerate infectivity in denatured PrP or create infectivity in synthetic or recombinant material. A number of candidate molecule classes have been considered, including other proteins, nucleic acids, glycosaminoglycans, sugars, and lipids. In general, the effectiveness of MALDIMS analysis of lipids is less predictable than for the analysis of peptides and proteins. Nevertheless, a combination of thin layer chromatography and MALDIMS was used to demonstrate that prion rods contain two host sphingolipids (41). Samples dried from chloroform with 5% methanol onto the MALDI target that had been pretreated with the matrix compound 2,5-dihydroxybenzoic acid gave normal mass spectra and fragment spectra using a technique known as post-source decay (PSD). In this way, it was established that chloroform/methanol extraction routinely yielded galactosylceramide and sphingomyelin, although in lower molar abundance than PrP. It is highly probable that these lipids were extracted from brain during the purification of PrPSc, which is known to be associated with membrane fractions, but it is not known whether they play any role in prion infectivity. The same group has recently identified the presence of an insoluble and inert polysaccharide scaffold in prion rods, remaining after prolonged protease digestion had removed at least 99.8% of PrP (42). The identification and analysis, which used direct chemical ionization, hydrolysis, conversion of the polysaccharide to alditol acetates and GCMS, showed predominantly 1,4-linked glucose structures. Such findings are relatively ubiquitous by this methodology; thus it is uncertain whether the presence of this polysaccharide in prions has any structural or functional significance. XI. CONCLUSIONS Mass spectrometry offers an array of powerful techniques for the analysis of the primary structure of proteins. By using a combination of

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these methods, PrPSc was confirmed to have an amino acid sequence identical to that predicted from the gene sequence. The nature of the posttranslational modifications was also delineated. Preliminary data on the heterogeneous N-linked sugars was substantiated by recent comprehensive studies that suggest that there are no unique components responsible for PrPSc formation. The GPI anchor was subjected to extensive analysis, revealing certain features never previously reported for comparable structures. However, further experiments suggested that these features are also present in PrPC and therefore do not provide the basis for an explanation of the different properties of PrPSc. Non-PrP peptides and other accessory molecules have been identified, but it appears unlikely that these are of any significance. The most difficult question that remains unanswered is whether a small fraction of molecules carrying a crucial modification has been overlooked. This is possible as one infectious prion unit is known to contain tens of thousands of PrP molecules. However, in summary, the weight of evidence from mass spectrometry supports the notion that the difference between PrPSc and PrPC underlying the infectivity of prions is not chemical but is essentially conformational (43). ACKNOWLEDGMENTS Mass spectrometry carried out in the UCSF Mass Spectrometry Facility was supported by NIH RR01614.

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12. Basler, K., Oesch, B., Scott, M., Westaway, D., Wälchli, M., Groth, D.F., McKinley, M.P., Prusiner, S.B., and Weissmann, C. (1986). Cell 46, 417–428. 13. Hope, J., Morton, L.J.D., Farquhar, C.F., Multhaup, G., Beyreuther, K., and Kimberlin, R. (1986). EMBO J. 5, 2591–2597. 14. Turk, E., Teplow, D.B., Hood, L.E., and Prusiner, S.B. (1988). Eur. J. Biochem. 176, 21–30. 15. Endo, T., Groth, D., Prusiner, S.B., and Kobata, A. (1989). Biochemistry 28, 8380–8388. 16. Haraguchi, T., Fisher, S., Olofsson, S., Endo, T., Groth, D., Tarentino, A., Borchelt, D.R., Teplow, D., Hood, L., Burlingame, A.L., Lycke, E., Kobata, A., and Prusiner, S.B. (1989). Arch. Biochem. Biophys. 274, 1–13. 17. Stahl, N., Borchelt, D.R., Hsiao, K., and Prusiner, S.B. (1987). Cell 51, 229–240. 18. Bolton, D.C., McKinley, M.P., and Prusiner, S.B. (1982). Science 218, 1309–1311. 19. Prusiner, S.B., Bolton, D.C., Groth, D.F., Bowman, K.A., Cochran, S.P., and McKinley, M.P. (1982). Biochemistry 21, 6942–6950. 20. McKinley, M.P., Bolton, D.C., and Prusiner, S.B. (1983). Cell 35, 57–62. 21. Pan, K.-M., Stahl, N., and Prusiner, S.B. (1992). Protein Sci. 1, 1343–1352. 22. Stahl, N., Baldwin, M.A., Teplow, D., Hood, L., Gibson, B.W., Burlingame, A.L., and Prusiner, S.B. (1993). Biochemistry 32, 1991–2002. 23. Baldwin, M.A., Falick, A.M., Gibson, B.W., Prusiner, S.B., Stahl, N., and Burlingame, A.L. (1990). J. Am. Soc. Mass Spectrom 1, 258–264. 24. Stahl, N., Baldwin, M.A., Burlingame, A.L., and Prusiner, S.B. (1990). Biochemistry 29, 8879–8884. 25. Ferguson, M.A.J., and Williams, A.F. (1988). Annu. Rev. Biochem. 57, 285–320. 26. Stimson, E., Hope, J., Chong, A. and Burlingame, A.L. (1999). Biochemistry 38, 4885–4895. 27. Rudd, P.M., Endo, T., Colominas, C., Groth, D., Wheeler, S.F., Harvey, D.J., Wormald, M.R., Serban, H., Prusiner, S.B., Kobata A., and Dwek, R.A. (1999). Proc. Natl. Acad. Sci. U.S.A. 96, 13044–13049. 28. Baldwin, M.A., Stahl, N., Reinders, L.G., Gibson, B.W., Prusiner, S.B., and Burlingame, A.L. (1990). Anal. Biochem. 191, 174–182. 29. Baldwin, M.A., Stahl, N., Burlingame, A.L., and Prusiner, S.B. (1990). In “Methods: A Companion to Methods in Enzymology” 1, 306–314. Academic Press, San Diego. 30. Stahl, N., Baldwin, M.A., Hecker, R., Pan, K.-M., Burlingame, A.L., and Prusiner, S.B. (1992). Biochemistry 31, 5043–5053. 31. Ferguson, M.A.J., Homans, S.W., Dwek, R.A., and Rademacher, T.W. (1988). Science 239, 753–759. 32. Baldwin, M.A., Wang, R., Pan, K.-M., Hecker, R., Stahl, N., Chait, B.T, and Prusiner, S.B. (1993). In “Techniques in Protein Chemistry” (R. Hogue Angeletti, ed.) Vol. IV, Academic Press, San Diego. pp. 41–45. 33. Harris, D.A., Huber, M.T., van Dijken, P., Shyng, S.L., Chait, B.T., and Wang, R. (1993). Biochemistry 32, 1009–1016. 34. Sandmeier, E., Hunziker, P., Kunz, B., Sack, R., and Christen. P. (1999). Biochem. Biophys. Res. Commun. 32, 1009–1016. 35. Weber, D.J., McFadden, P.N., and Caughey, B. (1998). Biochem. Biophys. Res. Commun. 246, 606–608. 36. Hornshaw, M.P., McDermott, J.R., and Candy, J.M. (1995). Biochem. Biophys. Res. Commun. 207, 621–629. 37. Hornshaw, M.P., McDermott, J.R., Candy, J.M., and Lakey, J.H. (1995). Biochem. Biophys. Res. Commun. 214, 993–999. 38. Loo, J.A. (1997). Mass Spectrom. Rev. 16, 1–23. 39. Last, A.M., and Robinson, C.V. (1999). Curr. Opin. Chem. Biol. 3, 564–570.

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40. Whittal, R.M., Ball, H.L., Cohen, F.E., Burlingame, A.L., Prusiner, S.B., and Baldwin, M.A. (2000). Protein Sci. 9, 332–343. 41. Klein, T.R., Kiersch, D., Kaufmann, R., and Riesner, D. (1998). Biol. Chem. 379, 655–666. 42. Appel, T.R., Dumpitak, C., Matthiesen, U., and Riesner, D. (1999). Biol. Chem. 380, 1295–1306. 43. Cohen, F.E., Pan, K.-M., Huang, Z., Baldwin, M.A., Fletterick, R.J., and Prusiner, S.B. (1994). Science 264, 530–531.

THREE-DIMENSIONAL STRUCTURES OF PRION PROTEINS BY KURT WÜTHRICH AND ROLAND RIEK Institut für Molekularbiologie und Biophysik, Eidgenössische Technische Hochschule Zürich, CH-8093 Zürich-Hönggerberg, Switzerland

I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Prions and Prion Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Natural and Recombinant Prion Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . C. NMR Information on Prion Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. About This Review . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. The NMR Structures of the Recombinant Bovine, Human, Mouse and Syrian Hamster Prion Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. A Global View of the Molecular Architecture . . . . . . . . . . . . . . . . . . . . . . . B. The Globular Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. The Flexible Tail . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Prion Protein Structure and the Species Barrier. . . . . . . . . . . . . . . . . . . . . . . . IV. Conclusions and Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

55 56 57 59 65 67 67 68 73 74 78 79

I. INTRODUCTION Transmissible spongiform encephalophathies (TSE) are neurodegenerative fatal diseases in mammalian species, which appear to occur as rare, “sporadic” disorders, but have also been traced to somatic mutations and to infectious transmission, including iatrogenic transfer (Prusiner, 1998; Weissmann, 1996). Scrapie in sheep, bovine spongiform encephalopathy (BSE) and the human TSEs kuru and CreutzfeldtJakob disease (CJD), and variants thereof have attracted interest far beyond the scientific community. The basic neurocytological lesions in TSEs are a progressive vacuolation of brain tissue, usually with extensive astroglial hyperthrophy and proliferation, which is sometimes described as “spongiform change” (Brown and Gajdusek, 1991). The “proteinonly hypothesis” (Alper et al., 1967; Griffith, 1967; Prusiner, 1982) suggests that TSEs are distinct from infectious diseases caused by bacteria, viruses, or viroids in that the origin of the disease is related to conformational alterations of an ubiquitous protein, (i.e., the “prion protein” [PrP]), and that nucleic acids are not essential for the propagation of the infectious agent. Accordingly, the normal “cellular form” of PrP (PrPC) is transformed into the disease-related, and possibly in itself infectious “scrapie form” (PrPSc) (Prusiner, 1998; Weissmann, 1996). 55 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved. 0065-3233/01 $35.00

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Not surprisingly, the observation that TSE pathology involves changes in the three-dimensional structure of a ubiquitous protein in healthy organisms has sparked keen interest in the molecular conformations of prion proteins. For PrPC from several mammalian species, atomic resolution nuclear magnetic resonance (NMR) structures are available, and these will be the major focus of this review. There may be a variety of disease-related forms of the prion protein, depending on factors such as the type and stage of the “prion disease” and the particular strain (Bessen et al., 1995; Safar et al., 1998). Nonetheless, it seems to be a common feature that PrPSc isolated from diseased brain tissue consists of prion protein aggregates, which have often been observed in the form of high molecular weight fibrils that have a significantly higher content of β-sheet secondary structure than PrPC (Prusiner, 1998). No atomic resolution three-dimensional structure has so far been reported for PrPSc. A. Prions and Prion Proteins According to Prusiner (1998) “the best current working definition of a prion is a proteinaceous infectious particle that lacks nucleic acid.” Historically, the concept that scrapie in sheep and other TSEs might be related to the presence of a particular protein rather than to nucleic acids dates back to experimental observations (Alper et al., 1967) and theoretical consideration (Griffith, 1967) in the 1960s. About a decade later it was demonstrated that a particular polypeptide sequence copurified with increasing TSE-infectivity in brain homogenates from laboratory animals in an advanced stage of TSE (Prusiner, 1982). From studies of this disease-related form of PrP, sufficient data on the amino acid sequence could be obtained to enable the cloning of a prion protein gene (named “Prnp” in the mouse) (Oesch et al., 1985). As a further key observation it was demonstrated that “knock-out” mice (i.e., mice with a nonfunctional Prnp gene) do not get sick on inoculation with TSE-infectious brain homogenate from other mice (Büeler et al., 1993). This indicates that host-encoded prion protein is required for the development of TSEs, and it was further shown that the PrPSc deposits in the brain of diseased individuals originate from hostencoded PrP. From these brief historical notes on the discovery of the prion protein we conclude that the benign PrPC form of the protein was isolated, purified, and further characterized without precise knowledge of its physiological role, although there are a variety of reports, sometimes apparently contradictory, on PrPC functions. Thus, in vivo and in vitro

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studies showed that PrPC binds copper (Brown and Besinger, 1998; Brown et al., 1997). There have also been observations that PrPC may be necessary for normal synaptic function (Collinge et al., 1994), and for the regulation of circadian activity rhythms and sleep (Tobler et al., 1996), but at least some mice strains are fully viable when devoid of functional Prnp genes (Büeler et al., 1992). Overall, no reliable information on the physiological role of the benign cellular form of the prion protein is available, and no common protein functions have been attributed to PrPC. B. Natural and Recombinant Prion Proteins The natural prion protein is encoded by a single exon as a polypeptide chain of about 250 to 260 amino acid residues, depending on the species (Oesch et al., 1985 Schätzl et al., 1995; Wopfner et al., 1999). Posttranslational modifications include cleavage of an N-terminal signal sequence of 22 residues, and a C-terminal signal sequence of about 23 residues, so that mature PrP consists of a single polypeptide chain with about 210 amino acid residues (Fig. 1) (Basler et al., 1986; Bazan et al., 1987). PrP contains a single disulfide bridge, as indicated in Figure 1 (Turk et al., 1988). PrPC also contains two glycosylation sites (Stahl and Prusiner, 1991), and, when isolated from natural sources, it was in some instances observed as a mixture of nonglycosylated, monoglycosylated, and diglycosylated protein (Rudd et al., 1999). PrPC contains numerous glycosylated variants (“glycoforms”), with at least 52 different bi-, tri-, and tetra-antennary N-linked oligosaccharides (Rudd et al., 1999). At the C-terminus, PrPC inserts into the cellular plasma membrane through a glycosyl-phosphatidyl-inositol (GPI) anchor (Stahl et al., 1987; Caughey and Raymond, 1991). For structural studies of PrPC, recombinant proteins were expressed in Escherichia coli. The constructs used contain either the intact polypeptide chain of the mature form of natural PrP (Fig. 1), possibly with some additional, construct-related residues at either chain end, or fragments thereof. Presently it appears impractical to envisage three-dimensional structure determinations with mammalian prion proteins from natural sources. However, sufficient amounts of natural PrPC have been isolated to enable qualitative comparative studies with the recombinant protein by optical spectroscopy. Overall, these experiments indicate close similarity between the natural and the corresponding recombinant prion protein. Thus, the circular dichroism (CD) spectrum of monomeric hamster PrPC extracted from hamster brains into a micellar environment of 30 mM n-octyl-β-glucopyranoside at pH 7.5 is typical for

58 FIG. 1. Amino acid sequence alignment of the human (h), bovine (b), mouse (m), and Syrian hamster (sh) prion proteins according to Schätzl et al. (1995). The data for the mature proteins, after cleavage of the signal sequences (see text), are shown. The complete sequence is given for hPrP, and for the other species only amino acid replacements, deletions, and insertions are explicitly indicated. The numeration at the top of the sequences corresponds to hPrP. For each protein the number in parentheses on the right indicates the numeration of the last residue in the constructs used for the NMR studies, where the numeration includes the N-terminal signal sequence. Below the sequences, black bars marked “O” represent “octapeptide repeats,” a gray bar “L” identifies an outstandingly hydrophobic and highly conserved peptide segment, which has been suggested to form a transmembrane helix (Hegde et al., 1998), and arrows mark the positions of the glycosylation sites and the C-terminal glyosylphosphatidylinositol (GPI) anchor. The disulfide bridge C179–C214 (numeration of hPrP) is schematically indicated.

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a protein with about 40% α-helix and little β-sheet content (Safar et al., 1993). Similarly, PrPC extracted from hamster brains into 10 mM aqueous sodium phosphate solution at pH 7.5 yielded CD and infrared spectra that are typical for α-helical proteins (Pan et al., 1993). Overall, the optical spectra thus indicate that natural PrPC, with its numerous posttranslational modifications has a similar conformation as recombinant PrP expressed in E. coli and dissolved in nondenaturing aqueous solvents. On a different line, evidence has also been presented that neither the C-terminal GPI-anchor nor the carbohydrate moieties in the positions 181 and 197 (Fig. 1) are essential molecular components with regard to the transformation of PrPC into forms that are associated with prion infectivity (Stahl et al., 1987). These data clearly emphasize the relevancy of structural studies with recombinant prion proteins for deeper insights into structure and function of natural prion proteins. C. NMR Information on Prion Proteins The intact polypeptide chain of recombinant PrP forms a rather unusual three-dimensional structure, which consists of two distinctly different chain segments of approximately equal size. The C-terminal half of the polypeptide chain forms a well-structured globular domain, whereas the N-terminal half forms a flexible extended “tail” (Fig. 2). This unusual global structure presents a quite convincing rationale for the fact that no suitable crystals for x-ray structure determination of prion proteins have so far been obtained. When using NMR spectroscopy for conformational characterization of PrPC, the structure of the C-terminal globular domain has been determined either as part of the intact polypeptide chain or in chain fragments that consist only of the domain. Even though the flexible tail does not adopt a unique, welldefined three-dimensional structure, informative NMR data can nonetheless be obtained. This section illustrates the procedures used for NMR studies of prion proteins by describing some details of the experiments with the mPrP, hPrP, and bPrP polypeptides. Similar techniques were used for the investigations with shPrP polypeptides (Donne et al., 1997; James et al., 1997; Liu et al., 1999). For three-dimensional structure determination by NMR in solution, one uses milligram quantities of homogeneous, pure protein, which is dissolved at millimolar concentration in 0.5 ml samples for the NMR measurements. The soluble form of the prion proteins discussed in this review was studied with an aqueous solvent of either 90% H2O/10% 2H O or 99.99% 2H O, which contained a suitable deuterated or other2 2 wise NMR-unobservable buffer, NaCl, and a trace of NaN3 to prevent

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Color Plates Legends FIG. 2. Cartoon of the three-dimensional structure of mPrP(23–231). Helices are yellow, β strands are cyan, segments with nonregular secondary structure within the C-terminal domain are green, and the flexibly disordered segments of residues 23–124 and 228–231 are represented by a succession of black lines representing virtual Cα–Cα bonds. The presentation is scaled to enable a comparison of the dimension of the globular domain with the length of the flexible “tails.” Some residue positions are identified with the sequence numbers, and limiting values for the rotational correlation times of 15N–1H groups in the globular domain and in the tails are also indicated. FIG. 3. Stereo view of the all heavy-atom presentation of a single conformer of the globular domain in bPrP(121–230). Brown color is used for 20 amino side chains that form a hydrophobic core of the protein molecule (see text). FIG. 5. Superposition of the mean NMR structures of the polypeptide segment 124–227 in bPrP(23–230) (violet) and bPrP(121–230) (green). A spline function was drawn through the Cα positions. The variable radius of the cylindrical rods is proportional to the mean global backbone displacement per residue, as evaluated after superposition for best fit of the backbone atoms N, Cα and C’ of the residues 125–227 in the two bundles of 20 energy-minimized conformers used to represent the two solution structures. FIG. 8. Cartoons of three-dimensional PrP structures. (A) Intact recombinant bovine prion protein, bPrP(23–230). (B) Intact recombinant human prion protein, hPrP(23–230). (C) Recombinant Syrian hamster prion protein, shPrP(29–231). The helices are colored green in (A), red in (B) and pink in (C); in all three structures the β strands are cyan, the segments with nonregular secondary structure within the globular domain are yellow, and the residues 23–120 in (A) and (B), and 29–124 in (C) of the flexibly disordered “tail” are schematically represented by yellow dots. FIG. 9. Pairwise comparisons of the mean NMR structure of the globular domain in bPrP(121–230) (green) with (A) hPrP(121–230) (red), (B) mPrP (121–231) (yellow), and (C) shPrP(121–231) (pink). Same presentation as in Figure 5: A spline function was drawn through the Cα positions. The variable radius of the cylindrical rods is proportional to the mean global displacement per residue, as evaluated after superposition for best fit of the atoms N, Cα, and C’ of the residues 124–227 in the 20 energy-minimized conformers used to represent the NMR structure. FIG. 10. NMR structure of mPrP(121–231) with indication of the hydrophobic core and the hydrogen bonds with amino acid side chains. The polypeptide backbone is represented by white ribbons and tubes. The hydrophobic core containing the residues 134, 137, 139, 141, 158, 161, 175, 176, 179, 180, 184, 198, 203, 205, 206, 209, 210, 213, 214, and 215 is shown in a yellow translucent envelope. In a shell surrounding the core, hydrogen bonds involving side chains (drawn as violet stick models) are represented by dashed cyan lines and labeled by a code of lower case letters that are referenced in the text. FIG. 11. Surface views of globular PrP domains with indication of the distribution of electrostatic charges. In the drawings on the left the orientation of the molecule is slightly changed relative to the “standard orientation” of Figs. 8 and 9, so that the residue 186 is approximately in the center of the front surface. The views on the right were generated from those on the left by a 180° rotation about a vertical axis. The electrostatic surface potential is indicated in red (negative charge), white (neutral), and blue (positive charge) coloring. (A) mPrP(121–231). (B) hPrP(121–230). (C) bPrP(121–230). FIG. 12. Mapping onto the NMR structure of mPrP(121–231) of those residue positions that are variable in different mammalian prion proteins (Fig. 1; Schätzl et al., 1995). A cartoon presentation of the polypeptide backbone is shown in white. The amino acid side chains in the variable positions are those of the mouse prion protein. Different colors for these side chains are used for easy reference in the text.

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bacterial growth. The protein construct, that is, the length of the polypeptide fragment and possibly designed amino acid exchanges in some sequence positions, and the solution conditions must be such that they prevent protein self-aggregation in the NMR sample. The expression system used must in most instances enable isotope labeling of the protein with 15N, 13C, and possibly 2H. For the production of isotope-labeled prion proteins, the initial studies with mPrP(121–231) relied on soluble periplasmic protein (Hornemann and Glockshuber, 1996). To obtain uniformly 15N-labeled and 13C, 15N-labeled mPrP(121–231), E. coli BL21 (DE3) cells (Studier and Moffat, 1986) harboring the T7-expression plasmid pPrP-C (Hornemann and Glockshuber, 1996) were used. Typically, to obtain about 20 to 30 mg of protein, cultures in 10 l of minimal medium containing as the sole nitrogen and carbon sources 15NH4Cl (2g/liter) and unlabeled glucose (5 g/liter), or 15NH4Cl (2 g/liter) and [13C6]-glucose (2 g/liter), respectively, were used. The cultures were induced with isopropyl-β-Dthiogalactoside, and purification to homogeneity started with soluble protein in the periplasmic fraction. mPrP(23–231) was initially prepared following a related strategy, where the purification started from protein in inclusion bodies to prevent proteolytic digestion of the tail (Hornemann et al., 1997). For the subsequent experiments with hPrP, bPrP, and mPrP, an alternative procedure was used with a T7-expression plasmid in E. coli that codes for a 17-amino acid N-terminal histidine tail with an engineered thrombin cleavage site (Zahn et al., 1996; 1997). The soluble protein fraction obtained after harvesting protein-rich inclusion bodies into an aqueous denaturant is added to nickel-nitrilotriacetic acid agarose resin, and the resin-bound protein is oxidatively refolded (Zahn et al., 1996) before thrombin cleavage and subsequent further purification to homogeneity. Cultures in 2 liters of minimal medium yield 20 to 40 mg of protein (Zahn et al., 1997). For stereospecific assignments of the methyl groups of Val and Leu, we decided to prepare prion protein with 10% “biosynthetically directed” 13C-labeling (Senn et al., 1989; Neri et al., 1989). To obtain concentrated protein solutions for NMR spectroscopy, Ultrafree-15 centrifugal filter devices (Millipore) have been used. NMR spectra are recorded using high-field spectrometers, in our laboratory with 1H frequencies of 500, 600, or 750 MHz. Conceptually different strategies yield information on the C-terminal globular domain and the N-terminal flexible tail (Fig. 2, see color insert). For the globular domain a standard protocol for NMR structure determination (Wüthrich, 1986; 1995) is used. For the sequential resonance assignment (Wüthrich, 1986; Wüthrich et al., 1982), triple-resonance experiments with the 13C, 15N-labeled proteins (Bax and Grzesiek, 1993) are

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primarily used, with supplementary data obtained from sequential and medium-range 1H–1H nuclear Overhauser effects (NOE) (Wüthrich, 1986). Based on the sequence-specific assignments, two-dimensional [1H,1H]-NOE spectroscopy (NOESY) (Anil-Kumar et al., 1980) and three-dimensional 15N- and 13C-resolved NOESY (Fesik and Zuiderweg, 1988) are used to collect an input of upper-limit constraints on 1H–1H distances (Wüthrich, 1986). Supplementary dihedral angle constraints are derived from measurements of scalar spin–spin coupling constants (Wüthrich, 1986) and from 13C chemical shifts (Spera and Bax, 1991; Wishart et al., 1995). The spectral analysis for obtaining resonance assignments, as well as for the collection of conformational constraints, is typically pursued with computer-supported interactive techniques, in our laboratory with the program XEASY (Bartels et al., 1995). With all the PrP polypeptides discussed in this review, nearly complete resonance assignments were obtained for the globular domain, which provided the basis for the collection of conformational constraints (Wüthrich, et al., 1982). As an illustration, the top two entries in Table I describe the input of conformational constraints for the structure calculation of the globular domain in bPrP. For the globular polypeptide segment there are approximately 15 NOE-derived upper distance constraints and 4 dihedral angle constraints per amino acid residue, which is representative for the globular domains of all prion proteins discussed here. Table I also presents data that enable an evaluation of the results of the structure calculation. One typically performs 100 calculations, starting from different random structures. The result of each individual calculation (Fig. 3, see color insert) can be evaluated by the residual violations of the experimental constraints (rows 3–9 in Table I). In our practice, the 20 “best conformers,” according to these criteria, are subjected to energy minimization (rows 10–13 in Table I), and the resulting group of energy-minimized conformers is used to represent the NMR structure. Visually, these 20 conformers represent a “bundle” of similar but not identical molecular models (Fig. 4A and B). The root mean square distances (RMSD) of these conformers to the mean atom coordinates represent the precision of the structure determination (rows 14–16 in Table I) (Wüthrich, 1986). The dispersion among the 20 conformers varies along the polypeptide chain, which may be quantitatively represented by “atom displacements” calculated for the individual amino acid residues (Billeter et al., 1989). An alternative to the bundle of Fig. 4A for the presentation of the polypeptide backbone is a spline function through the Cα positions, with the radius of the cylindrical rod representing the variable average atom displacements calculated for the individual residues along the polypeptide

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TABLE I NMR Structure Calculation of the Globular Domain in bPrP Quantity NOE upper distance limitsb Dihedral angle constraintsb Residual target function (Å2)c Residual NOE violationsd Number ≥ 0.1 Å Maximum (Å) Residual torsion angle violationsd Number ≥ 2.0 deg. Maximum (deg.) AMBER energies (kcal/mol)d Total Van der Waals Electrostatic RMSD to mean coordinates (Å)e N, Cα, C′ (125–227) All heavy atoms (125–227)

bPrP(23–230)a

bPrP(121–230)a

1,576 469 1.15 ± 0.28

1,797 483 0.79 ± 0.19

30 ± 5 0.15 ± 0.01

20 ± 3 0.14 ± 0.01

0.4 ± 0.5 2.4 ± 1.1

0.3 ± 0.4 2.1 ± 0.7

–4,892 ± 86 –316 ± 14 –5,208 ± 84

–4,952 ± 78 –344 ± 9 –5,296 ± 76

0.78 ± 0.12 1.21 ± 0.11

0.69 ± 0.11 1.10 ± 0.10

The structure of the globular domain was computed independently from NMR data collected with intact recombinant bPrP, bPrP(23–230), and with the polypeptide fragment bPrP(121–230). The two structures are characterized by a bundle of 20 conformers each, which were obtained by repeated structure calculations starting from different random structures. Except for the top two rows the numbers in the table are the mean values and the standard deviations for the 20 conformers. b These two rows indicate the total number of conformational constraints used as input for the structure calculations. c Residual value of the DYANA target function (Güntert et al., 1997) before energy minimization; for an individual calculation a small value indicates good convergence. d Parameters that characterize the quality of the structure calculation after energy minimization. e The root mean square deviations (RMSD) describe the global precision of the structure determination. a

chain (Fig. 5, see color insert). In our laboratory we use the programs DYANA (Güntert et al., 1997) for the structure calculations, OPALp (Luginbühl et al., 1996; Koradi et al., 2000) for energy minimization, and MOLMOL (Koradi et al., 1996) for analysis and visual presentation of the results of the structure calculations. Measurements of 15N{1H}-NOEs, and longitudinal and transverse 15N spin relaxation times can provide supplementary information on the globular PrP domain in the form of data on the internal mobility along the polypeptide chain (Peng and Wagner, 1992). Furthermore, realtime

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FIG. 4. Stereo views of the globular domain of bPrP. (A) Polypeptide backbone superposition of the 20 conformers used to represent the NMR structure of the globular domain in bPrP(121–231). (B) Same as (A), except that all heavy atoms are drawn. The same orientation of the molecule was used in the Figures 3, 4A and 4B.

measurements of the exchange of amide protons with 2H on dissolving the protein in 2H2O solution yield information on the solvent accessibility of the amide groups, as expressed by “protection factors” (Bai et al., 1993). The protection factors manifest conformational equilibria between a manyfold of protein states that contain individual amide groups in solvent-accessible or solvent-inaccessible locations, respectively. For the N-terminal tail (Fig. 2), small dispersion of the 1H chemical shifts and small amide proton protection factors showed early on that this polypeptide segment is predominantly in an extended “random coil” state (Donne et al., 1997; Riek et al., 1997). Sequential resonance assignments for the tail can be obtained with triple resonance experiments optimized for this special situation (Liu et al., 2000), and with the exception of the octapeptide repeats (Fig. 1), nearly complete assignments have been obtained. This provides a basis for the analysis of NMR data such as heteronuclear 15N{1H}-NOEs and 15N relaxation times, which

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provide information on the frequency of intramolecular rate processes (Peng and Wagner 1992) also when no unique three-dimensional structure can be defined in “unfolded” polypeptide chains (Fig. 2). Some key data for the characterization of the PrP tail are displayed in Figs. 6 and 7. Figure 6A shows a [15N, 1H]-correlation experiment (COSY), where each 15N–1H moiety is represented by a cross-peak that connects the 15N chemical shift (along the vertical frequency axis, ω1) with the 1H shift of the directly bound hydrogen atom (along the horizontal frequency axis, ω2). This experiment thus provides a “NMR fingerprint” of the protein. In a different experiment one measures nuclear Overhauser effects (NOE) between directly bound 15N and 1H spins (15N{1H}-NOE). Rapidly moving 15N–1H groups in flexible polypeptide segments have negative values for the 15N{1H}-NOEs, whereas positive values are seen for “immobilized” 15N–1H groups in globular proteins (Wüthrich, 1986). In other words, positive intensities of the 15N{1H}-NOEs correspond to 15N–1H groups with rotational correlation times, τc, longer than about 2 ns, and negative intensities are seen for correlation times shorter than approximately 0.5 ns. Figure 7 then shows that in bPrP(23–231) the tails from residues 23–121 and 228–230 are highly flexible, whereas the globular domain with residues 122–227 shows positive NOEs that indicate restricted mobility, as is typical for globular proteins (see also Fig. 2). The 15N{1H}-NOEs can also be presented in a two-dimensional spectrum, using the [15N,1H]-COSY-relayed 15N{1H}-NOE experiment. Positive and negative cross-peaks may then be displayed in separate subspectra (Fig. 6, B and C), which affords a nice illustration of the different dispersion of the 1H chemical shifts in the unfolded and folded parts of the protein (Wüthrich, 1986). The spectrum of Fig. 6B is typical for a globular domain, with the backbone amide proton chemical shifts spread out from 7 to 10 ppm, and the spectrum of the tail in Fig. 6C contains backbone amide proton peaks only between 8.0 and 8.7 ppm, which is typical for an extended “random coil” polypeptide chain. D. About This Review Until the determination of the NMR structure of mPrP(121–231) in 1996 (Riek et al., 1996), knowledge of the three-dimensional structure of PrPC was based on optical spectroscopy data (Pan et al., 1993; Safar et al., 1993), a wealth of biochemical observations, and physiochemical studies with synthetic polypeptide fragments of PrP that also included NMR investigations (e.g., Zhang et al., 1995; see also Tagliavini et al., 2001). Given the scarcity of experimental data, for PrPC as well as for PrPSc, much effort was devoted to three-dimensional molecular model-

FIG. 6. Two-dimensional NMR spectra of bPrP(23–230) (protein concentration 1 mM, solvent 90% H2O/10%D2O, 10 mM sodium acetate, pH = 4.5, T = 20°C, 1H frequency = 500 MHz). (A) [15N, 1H]-correlation spectrum (COSY). Each 15N–1H moiety in the protein is represented by a cross-peak. (B) Subspectrum containing the positive peaks from a [15N,1H]-COSY-relayed 15N{1H}-NOE experiment, representing crosspeaks of 15N–1H groups in the globular domain of bPrP(23–230). (C) Subspectrum containing the negative peaks from the same experiment as in (B), representing 15N–1H groups in flexibly disordered regions. The rectangle in (C) surrounds the spectral region of the side chain amide and guanidinium resonances. The circle contains the tryptophan indole resonances.

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FIG. 7. Plot versus the amino acid sequence of the relative intensities, Irel, of the steady-state 15N{1H}-NOEs of the backbone amide groups of bPrP(23–230). In the box from positions 51 to 91 the pattern of NOE values is indicated for the octapeptide insertion between the positions 67 and 68 (Fig. 1), where the open rectangle indicates that only one value was measured for two overlapped Gly residues. The circles indicate that identical patterns prevail for the other octapeptide repeats (see text). The regular secondary structures of the globular domain are indicated at the bottom, and the first residue with a positive NOE-value, Val122, is identified.

ing. A heuristic approach was used, which included prediction of regular secondary structures from the amino acid sequence, evaluation of favorable packing of regular secondary structure elements, and consideration of selected in vitro and in vivo experimental data (Huang et al., 1994, 1996). In addition to structure predictions for PrPC and PrPSc, these modeling efforts also led to an outline for a hypothetical pathway of the disease-related transition from PrPC to PrPSc. This work has been extensively reviewed elsewhere (e.g., Cohen, 1999; Prusiner, 1996), and the investigations with peptide fragments from PrP are covered elsewhere in this volume (Tagliavini et al., 2001). We therefore decided to focus this review entirely on the atomic resolution structures of soluble recombinant mammalian prion proteins, and on implications of these three-dimensional structures for possible physiological roles of PrPC, including functional features that might be relevant for the pathology of TSEs in humans and other mammalian species. II. THE NMR STRUCTURES OF THE RECOMBINANT BOVINE, HUMAN, MOUSE, AND SYRIAN HAMSTER PRION PROTEINS A. A Global View of the Molecular Architecture The previously introduced architecture of the intact mature polypeptide chain of mPrP (Fig. 2) is common to all four presently available PrP

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structures (Fig. 8, see color insert). The N-terminal domain with residues 23–124 forms a flexibly extended “tail”, which is an unusual feature compared with the presently available protein structure database. The C-terminal domain has the shape of a flat ellipsoid and contains common α-helical and β-sheet regular secondary structures. For the individual species, complete structure determinations of the globular domain have been performed with different polypeptides, which vary in the length of the tail (for details see the following section B). In mPrP and bPrP there is no measurable impact of the tail length on the globular domain, but in hPrP the full length tail appears to measurably affect the C-terminal turns of the helices 2 and 3 (see Section II. B. 3. below). B. The Globular Domain This section starts with a detailed description of the globular domain in mPrP, which is the first prion protein for which a three-dimensional structure was obtained (Riek et al., 1996). Considering the high homology on the levels of both sequence (Fig. 1) and three-dimensional architecture (Fig. 8), the description of the proteins from the other three species concentrates on the structural differences relative to mPrP, and on data that were collected only for individual species. 1. Murine PrP The structure of the globular domain of mPrP has been calculated using data collected with mPrP(121–231) (Riek et al., 1998; Calzolai et al., 2000) and mPrP(23–231) (Riek, 1998). In both proteins, the RMSD values for the backbone and for all heavy atoms, respectively, of the residues 125–226 are 0.8 Å and 1.4 Å. The global polypeptide fold of the mPrP globular domain (Fig. 2) contains three α helices with residues 144–154, 175–194 and 200–226, and two β strands with residues 128–131 and 161–164. The thickness of the yellow cylindrical rod in Fig. 9B (see color insert) represents the variable precision of the backbone structure determination in mPrP(121–231) along the sequence from residues 124–227. Strongly increased disorder is seen for the C-terminal two turns of helix 3, which also diverge from the helix axis defined by the residues 200–218, and a loop connecting the second β strand with helix 2, approximately from residues 166 to 172. The poor definition of this loop is related to the fact that the backbone 15N–1H groups of the residues 167–171 were not observed in the NMR spectra, presumably because of line broadening due to slow conformational exchange (Riek et al., 1998). Best defined are the core parts of the regular secondary

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structures, whereas the connection to the flexible tail at residue 124, the connecting segment from the first β strand to helix 1, and the last two turns of helix 2 are somewhat less well defined. In the β sheet the residues 129, 131, 161, 163, and 164 show typical β-sheet deviations of the 13Cα chemical shifts from the random coil values (Spera and Bax, 1991), but the Hαchemical shifts and the 3JHNα coupling constants in the first β strand are close to the random coil values, which may indicate local conformational averaging. There is also a hydrogen bond from the amide proton of Met134 to the carbonyl oxygen of Asn159 that would be compatible with an elongation of the β sheet toward the first helix, with a β bulge at residue 132. These observations are of interest, as they are indicative of a certain degree of “dynamic plasticity” of the β sheet. The structure of mPrP contains a tightly packed hydrophobic core formed by 20 amino acid side chains. In Fig. 10 the outer confines of the core are indicated by a translucent sheet. 13 residues in the hydrophobic core come from the helices 2 and 3, which are covalently linked by the disulfide bridge Cys179–Cys214 and have further mutual interactions through the residues Phe175, Val176, Val180, Ile184, Val203, Met206, Val210, and Val215. The side chains of Met205, Val209, and Met213 in the helix 3 interact with Met134, Pro137, Ile139, and Phe141 in the loop between the first β strand and the first helix (Fig. 2). Pro 158 and Val161 are adjacent to and within the β sheet, respectively, and Phe198 is located in the loop between the helices 2 and 3. Among the 20 core residues, 16 positions contain identical amino acids in the four species of Fig. 1, and the remaining 4 positions have exclusively conservative variations, or are exchanged pairwise in a correlated fashion so that the local packing arrangement is conserved (Riek et al., 1998). These residues are highly conserved also in a wider array of mammalian PrPs (Schätzl et al., 1995; Wopfner et al., 1999), indicating that the core structure of the globular domain seen in mPrP may be maintained in all mammalian species. The hydrogen bonds in the β sheet have been described in the preceding section, and with the exception of the C-terminal turns, which may have some 310-helix character, regular backbone–backbone α-helix hydrogen bonds are present in the three helices. In addition, the hydrophobic core is surrounded by an “outer shell” of protein structure that contains 18 hydrogen bonds with amino acid side chains (Fig. 10, see color insert), which include some medium-range interactions (Wüthrich, 1986) within the three helices, and a variety of longer-range interactions. In the helices the nonstandard hydrogen bonds include Asn143Hδ–OεGlu146 at the N-terminus of helix 1 (b in Fig. 10),

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Asn171Oε–Hδ Asn174 at the start of helix 2 (I in Fig. 10), Cys179O’– HδThr183 (m in Fig. 10), and a capping box of Thr199HN–Oδ Asp202 and Asp202HN–OγThr199 at the N-terminus of helix 3, which is further stabilized by a side chain–side chain hydrogen bond Thr199OHγ–OδAsp202 (r, s and t in Fig. 10). Among the longer-range interactions that connect different groups within the outer shell, there are the two hydrogen bonds Tyr128Hη–OδAsp178 and Tyr162HN–OγThr183 (a and j in Fig. 10), and the salt bridge Arg164–Asp178 (k in Fig. 10), which hold the β sheet against helix 2. A salt bridge Arg156–Glu196 (g in Fig. 10) orients the C-terminal end of helix 1 toward the loop connecting the other two helices. Similar to the aforementionned hydrophobic core residues, most of the hydrogenbonded amino acid side chains are strictly conserved (Fig. 1) (Schätzl et al., 1995; Wopfner et al., 1999). Nonconserved hydrogen bond-forming side chains include Asn143, Arg164, and Asn174; but these are all exchanged by amino acid types that enable maintenance of corresponding hydrogen bonds in the prion proteins from different mammalian species. As mentioned before, the mPrP globular domain has the shape of a flat ellipsoid. Its surface is characterized by a pronouncedly uneven distribution of positively and negatively charged residues between the two flat surfaces (Fig. 11A, see color insert). Considering that in a membrane-associated state the positively charged surface would be expected to be preferentially oriented toward the lipid phase, it is worth noting that the two glycosylation sites at Asn181 and Asn197 are located on the negatively charged surface of the protein, which would then presumably be solvent-exposed on the cell surface (Riek et al., 1996). Inspection of the amino acid sequences corresponding to the globular domain in mammalian species (Fig. 1; Schätzl et al., 1995; Wopfner et al., 1999) indicates a high degree of conservation, which should be sufficient for maintaining the same molecular architecture throughout. Nonetheless, there are amino acid replacements that may be of functional interest. Figure 12 (see color insert) affords a survey of the distribution of these variable sites in the three-dimensional structure of mPrP. Most strikingly, the relatively large number of species variations in the peptide segments 166–170 and 215–230 (Fig. 1) are all found in a continuous area of the protein surface (blue in Fig. 12). As was previously pointed out by Billeter et al. (1997), the propensity for intermolecular hydrogen bond formation in this surface area changes from species to species as a result of nonconservative exchange of polar and charged amino acids (Fig. 1). On the opposite surface of the protein, three nonconservative amino acid exchanges are located on helix 1 and

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the adjacent loops (green in Fig. 12), and a local variation in the electrostatic charge occurs on the surface of helix 2 at residue 186 (orange in Fig. 12). Additional amino acid replacements located in the interior of the molecule (violet in Fig. 12) are conservative exchanges of different hydrophobic side chains. 2. Syrian Hamster PrP The structure of the globular domain of shPrP was computed from data collected with shPrP(90–231) (James et al., 1997; Liu et al., 1999). Its global architecture coincides with that of mPrP (Fig. 8C), but there are two striking differences in local structure. In contrast to mPrP, all the resonances in the polypeptide segment of residues 167–175 were observed in shPrP(90–231), and the loop between the second β strand and helix 2 is well defined (pink structure in Fig. 9C). Furthermore, helix 3 is a well-defined, straight α helix up to residue 227 (Fig. 8C; pink structure in Fig. 9C). Otherwise, the precision of the structure determination varies almost identically along the polypeptide chain in mPrP and shPrP (Fig. 9, yellow structure in B and pink structure in C). The aforementionned indications of a certain degree of “conformational plasticity” of the β sheet in mPrP have also been noticed in shPrP(90–231) (Liu et al., 1999). The polypeptide segment connecting the first β strand with helix 1 is arranged slightly differently in mPrP and shPrP, as can readily be seen from the superpositions with bPrP in Fig. 9B and C. This is probably related to the species variation of the amino acid in position 139 (Fig. 1). 3. Human PrP The globular domain structure, which has the same architecture as mPrP, was independently determined from data collected with hPrP(23–230), hPrP(90–230), and hPrP(121–230) (Zahn et al., 2000). In the intact protein, the regular secondary structure elements coincide identically with those in hPrP(90–230) and hPrP(121–230), with the residues 128–131 forming the β strand 1, 144–154 the α helix 1, 161–164 the β strand 2, 173–194 the α helix 2, and 200–228 the α helix 3. A striking local difference relative to mPrP is seen for helix 3, which is more regular and better defined for the segment of residues 220–228 (Fig. 8B). As in mPrP(121–231), there is a hydrogen bond from the amide proton of Met134 to the carbonyl oxygen of Asn159 in all three hPrP polypeptides, which is reminiscent of an irregular, β-bulge-type elongation of the β sheet toward the helix 1. In the three hPrP polypeptides the globular domain has nearly identical side chain conformations of the interior hydophobic amino acids. Overall, the regular

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secondary structures are well defined (Fig. 9A, red structure), but similar to mPrP several backbone resonances of the residues 166–172 were not observed in the NMR spectra, and as a result this loop is less well defined. Somewhat decreased precision of the structure determination is seen also at the end of helix 2 and the following loop, and for the last two turns of helix 3. In the investigations of hPrP, special efforts have been made to characterize conformational equilibria in the less precisely structured polypeptide segments by studies of the 13Cα chemical shifts and measurements of amide proton protection factors (Hosszu et al., 1999; Zahn et al., 2000). In contrast to the NOE intensity, which depends on the inverse sixth power of the 1H–1H distance and therefore usually manifests only the folded form with the shortest distance (Wüthrich, 1986), the differences between observed and random coil 13Cα chemical shifts, ∆δ(13Cα), can be qualitatively related to the population of regular secondary structures (Spera and Bax, 1991; Wishart and Sykes, 1994). For all 13Cα atoms located within the α helices of hPrP, the resonances show the expected downfield shifts relative to the random coil values, but smaller values of ∆δ(13Cα) indicate for the C-terminal two turns of each of the helices 2 and 3 that the α-helical structure is in equilibrium with unfolded forms of the polypeptide. This was corroborated by the observation that all amide protons in the hydrogen bonds of the helices 2 and 3 were measurably protected against exchange, except for the residues 187–194 in helix 2, and the residues 225–228 in helix 3 (Zahn et al., 2000). The 13Cα chemical shifts further provided indications for transient intramolecular contacts between the flexibly disordered tail of hPrP and the globular domain. Small 13Cα chemical shift differences between hPrP(23–230) and hPrP(121–230) were observed for the residues 187–193 in the C-terminal part of helix 2, and for the residues 219–226 in the C-terminal part of helix 3. [Differences in 13Cα chemical shifts had also been found for the residues 187–193 between shPrP(29–231) and shPrP(90–231) (Donne et al., 1997), but were subsequently attributed to small pH differences between the two protein samples (Liu et al., 1999)]. In hPrP the ends of the helices 2 and 3 appear to be slightly stabilized in the presence of the intact tail, with the implicated increase of helix population amounting to a few percent, but the mean structure is not visibly affected by the different tail lengths. Since an effect of the tail length on helix 3 appears so far to be unique for hPrP(23–230), it was tentatively attributed to the high negative charge located near the end of helix 3 in hPrP (Zahn et al., 2000).

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4. Bovine PrP The structure of the globular domain was determined from data collected with the polypeptides bPrP(23–230) and bPrP(121–230) (LópezGarcía et al., 2000), as described in Section I. C. The domain structure of bPrP(23–230) is indistinguishable from that of bPrP(121–230) (Fig. 5). It is also essentially identical to hPrP (Fig. 9A), but the backbone fold shows larger local differences relative to mPrP (Fig. 9B). On the other hand, the packing of the hydrophobic core side chains (Fig. 3) coincides closely with mPrP (Fig. 11). bPrP coincides with hPrP and mPrP in that the backbone amide protons of Asp167, Ser170, Asn171 and Phe175 were not observed in the NMR spectra. Similar data on conformational equilibria and intermolecular rate processes were obtained for bPrP (Figs. 6 and 7) and hPrP. C. The Flexible Tail As described in Section I. C, NMR experiments can provide a characterization of polypeptide conformation even when the chain is flexibly disordered and does not adopt a discrete set of three-dimensional structures. Thus, based on data corresponding to those for bPrP in Figures 6 and 7 (López-García et al., 2000), it was demonstrated that the N-terminal half of the intact polypeptide chain forms a flexible tail in mPrP (Riek et al., 1997), shPrP (Donne et al., 1997), and hPrP (Zahn et al., 2000). In the initial study of mPrP(23–231), all residues in the segment 23–120 were treated as one “class,” with the common property of being flexible on a subnanosecond time scale (Riek et al., 1997). Sequence-specific resonance assignments have then been obtained for shPrP(29–231) (Donne et al., 1997) and subsequently also for mPrP(23–231) (Riek, 1998), hPrP(23–230) (Liu et al., 2000), and bPrP(23–230) (Lopez et al., 2000). Figure 7 illustrates the extent of the assignments that could be achieved. In all four species these are nearly complete for the residues 23–50 and 92–120. For the octapeptide repeats from positions 51 to 91 (Fig. 1), individual sequence-specific assignments were obtained only for a small number of residues in the first and last repeats, as chemical shift degeneracy prevented further assignments of individual residues. Experience with NMR experiments specifically adapted for use with flexible polypeptide chains indicates that these residual degeneracies will not be resolved readily (Liu et al., 2000). The extensive chemical shift degeneracy of corresponding residues in the different octapeptide repeats clearly corroborates the “random coil” behavior of the tail.

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Special interest has focused on the potential transmembrane segment that immediately precedes the globular domain (L in Fig. 1). In shPrP(90–231), evidence has been presented for some preferred structure of this peptide segment, which might also involve interactions with the residues 125–128 (Liu et al., 1999). Numerous studies used synthetic polypeptides that include this hydrophobic segment (L in Fig. 1) (see Tagliavini et al., 2001). Another focus is the role of the tail in copper binding by PrP (e.g., Brown et al., 1997). A “plausible model” for a Cu2+ complex with the tail segment 58–91 of shPrP was advanced based on studies of Cu2+ binding to synthetic polypeptides consisting of parts or all of this sequence (Viles et al., 1999): a circular array of –His61–Cu2+–His69–Cu2+–His77–Cu2+–His85– Cu2+– forms the core of a multilooped structure, with outer loops formed by the polypeptide segments 62–68, 70–76 and 78–84. Based on the sequence-specific resonance assignments, further detailed structural information on the PrP tail can be anticipated. Overall, structural data on the tail that have been collected with the intact proteins are still scarce, but work is in progress in our laboratory as well as elsewhere. III. PRION PROTEIN STRUCTURE AND THE SPECIES BARRIER The “species barrier” for TSEs describes the fact that although transmission between different individuals of the same mammalian species occurs efficiently, the process of infection between different species may be either inefficient or completely absent (Prusiner, 1998; Weissmann, 1996). If one accepts the protein-only hypothesis (Alper et al., 1967; Griffith, 1967; Prusiner, 1982) so as to attribute a key role to PrP in the TSE-infection process, either as the infective agent or in a supporting role, one will quite necessarily infer that the stringency of the species barrier should be related to the degree of structural homology between the prion proteins of the host and the infectious material (Scott et al., 1989; 1993). This issue has been extensively discussed on the basis of a sizeable database of PrP sequences (Schätzl et al., 1995; Wopfner et al., 1999). Deeper insight may be gained with the additional availability of three-dimensional structures, because amino acid exchanges between different species may affect the structure of PrPC and/or the structure of PrPSc, as well as interactions of either or both forms of PrP with species-specific receptor sites. In this chapter we investigate possible correlations between species variations of the PrPC three-dimensional structure and the stringency of the species barrier between different pairs of mammalian species.

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Pairwise superpositions of the four presently known PrPC structures (Fig. 9) document the previously mentioned close similarity of the globular domains in bPrP, hPrP, mPrP, and shPrP; hPrP is closest to bPrP (Fig. 9A), with an RMSD value of 0.98 Å between the mean structures. The corresponding values for the superpositions of bPrP with mPrP and shPrP (Fig. 9B and C) are 1.66 Å and 1.68 Å, respectively, where local differences between the backbone conformations are mainly seen for helix 1, the loop of residues 166–172, and the end of helix 3. The helix 1 overlaps closely between bPrP and hPrP (Fig. 9A), whereas for mPrP and shPrP, it is displaced toward the core of the protein compared with bPrP (Fig. 9B and C). The helix 3 in bPrP, hPrP, and shPrP is a regular α helix up to approximately residue 226 (Fig. 8), whereas in mPrP there are deviations from a regular α-helical structure and significantly increased structural disorder from residue 219 onward (Fig. 9B). The loop 166–172 is fully NMR-observable only in shPrP (Liu et al., 1999). The loop 166–172 and the helix 3 have been recognized as “conformational markers” that are located near the protein surface and are therefore defined with only limited precision in the NMR structure (Calzolai et al., 2000). The two opposite molecular surface regions formed by helix 1, and by the loop 166–172 and helix 3, respectively (Fig. 9), have been proposed, on the basis of experiments with transgenic mice and hamsters, as sites for intermolecular interactions that might contribute to the species barrier (Prusiner, 1998; Telling et al., 1994; 1995). For helix 1, a closer inspection shows that in addition to the near-identity of the backbone structure in bPrP and hPrP (Fig. 9A), the two species have identical sequences in helix 1 and the adjoining loops (Fig. 1). In contrast, the three sequence positions 143, 145, and 155 in or immediately adjacent to helix 1 show nonconservative amino acid exchanges between bPrP (or hPrP) and either mPrP or shPrP (Fig. 1). These variations include Ser/Asn, Tyr/Trp, and His/Tyr or Asn substitutions on the protein surface, which can be expected to modify the propensity for intermolecular interactions with the protein surface near helix 1. In additional studies, the conformational markers in the surface area formed by helix 3 and the loop 167–172 (Fig. 2) were related to singleamino acid exchanges in hPrP (Calzolai et al., 2000). In three variant proteins the residues Met166 and Arg220 of hPrP were replaced with Val and Lys, respectively, which are the corresponding residues in mPrP, and Ser170 in hPrP was replaced with Asn, which is the corresponding amino acid in shPrP. Table II summarizes the results of complete NMR structure determinations of these three hPrP analogs, which show variations in the two conformational markers within the scaffold

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TABLE II Conformational Markers in Mammalian Prion Proteins Proteina hPrP(121–230) mPrP(121–231) bPrP(121–230) shPrP(90–231) hPrP(M166V) hPrP(S170N) hPrP(R220K)

166–172, 175b

Helix 3c

+

+ (0.49/–) (0.68/1.59) + (0.38/0.71) + (0.33/0.65) + (0.49/0.91) + (0.49/0.44) (0.62/1.12)

+

In hPrP(M166V) and hPrP(R220K) the amino acid of wildtype hPrP is replaced by the corresponding amino acid in mPrP. In hPrP(S170N), the exchange is between wild-type hPrP and shPrP. b A “+” sign indicates that [15N,1H]-COSY cross-peaks were observed and assigned for the entire loop 166–172 and for residue 175. c A “+” sign indicates that for the residues 200–226 the RMSD to the mean coordinates for N, Cα and C′ among the 20 conformers used to describe the NMR structure is < 0.5 Å, and that the RMSD of the mean structure relative to the mean hPrP(121–230) structure is < 1.0 Å (the two RMSD values are given in parentheses). a

of the preserved global structure. The helix 3 is as well defined in hPrP(S170N) as in hPrP, and for the helix 3 the differences between these two proteins fall within the conformation space spanned by the bundles of 20 conformers used to characterize each of the two NMR structures (Table II). In hPrP(M166V), the helix 3 is well defined up to residue 226 but shows a slight deviation from the straight helix axis after residue 220, which is manifested in an increase of the local RMSD value for the residues 200–226 in hPrP(M166V) compared with wildtype hPrP (Table II). In hPrP(R220K) the helix 3 is well characterized up to residue 219, but then it is less well ordered and shows a more pronounced deviation from the straight helix axis than hPrP(M166V). This is again clearly manifested in an increase of the RMSD value for residues 200–226 (Table II). For the second conformational marker there is a clear-cut, qualitative difference among the three variant proteins. In hPrP(R220K) and hPrP(M166V) the [15N,1H]-COSY crosspeaks for the residues 167, 169, 170, 171, and 175 were not observed, which is similar to previous observations in hPrP, mPrP, and bPrP, whereas in hPrP(S170N) all the cross-peaks in the polypeptide segment

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166–175 have been detected, which coincides with corresponding observations in shPrP (Liu et al., 1999). Overall, these data on the impact of single-amino acid replacements on static and dynamic aspects of the hPrP conformation (Table II) not only establish links between sequence and conformation, but support further that the independently observed local conformational differences in the surface area of helix 3 and the loop 166–172 between hPrP, bPrP, mPrP, and shPrP (Table II) are significant, although they are located in a poorly ordered molecular region (Fig. 9). Evidence for an involvement of the molecular surface area of the blue side chains in Fig. 12 in species-specific intermolecular interactions includes that the species barrier for transmissible mink encephalopathy between ferret and mink must relate to the only two residue variations between the two species, which are in the positions 175 and 220 (Bartz et al., 1994), and that the species barrier between man and rabbit has been related to the replacement of Asn174 in hPrP by Ser (Loftus and Rogers, 1997). Furthermore, from experiments with transgenic mice a discontinuous epitope of residues 168, 172, 215, and 219 (numeration for hPrP) has been suggested to be involved in binding of PrPC to a conversion factor, “protein X,” which would mediate the transformation of PrPC to PrPSc (Kaneko et al., 1997; Telling et al., 1995), and the fragment 225–231 in mPrP represents an epitope for a monoclonal antibody (R2) that strongly reacts with mPrP and shPrP, but not with hPrP and bPrP (Williamson et al., 1998). On this background the close coincidence between the globular domains of hPrP and bPrP is quite intriguing. If the conformation of the surface area of PrPC formed by helix 1 is a factor that contributes to species barriers, this contribution to the barrier between cattle and humans will be small or completely absent, whereas it could have an important influence on the barrier between either of these two species and the mouse or the Syrian hamster. Similar considerations for the conformation of the PrPC surface area formed by the loop 166–172 and the C-terminal part of helix 3 indicate that contributions to a species barrier between humans and cattle would be small, whereas conformational differences might contribute to or even represent a dominant factor in the species barriers between cattle (or humans) and mouse or Syrian hamster (Collinge et al., 1995; Telling et al., 1994; 1995), as well as between mouse and Syrian hamster (Scott et al., 1989; 1993). Variation of the electrostatic charge distribution on an otherwise invariant molecular surface might influence intermolecular recognition of PrP. Although the uneven charge distribution between the two flat surfaces is preserved among different PrPC structures (Fig. 11),

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there are nonetheless distinct differences. Thus, hPrP is unique among the species of Fig. 1 in carrying a charge of –3 in the surface region formed by helix 3 and the loop of residues 166–172, with the corresponding charge in the other three species being –1. Similarly, bPrP is unique in carrying a negative charge in position 186, which contains an uncharged, polar residue in the other three species (Fig. 1). The resulting different surface charge distributions in bPrP and hPrP are readily apparent in Figure 11B and C, which also shows that there are no charge variations between these two species outside of the loop 166–172, position 186, and the C-terminal half of helix 3 (see also Fig. 1). Overall, it has been concluded from these comparative studies that surface charge variation is the sole potentially relevant feature of the PrPC structure with regard to a species barrier between humans and cattle (Lopez et al., 2000). In contrast, variation of the surface conformation would appear to be an additional factor that might affect species barriers between humans or cattle and the laboratory animals of Figure 1. IV. CONCLUSIONS AND OUTLOOK The prevalence of identical global architectures for the four proteins of Figure 1 (Figs. 2, 8, and 9) implies that conformational transitions between the ubiquitous cellular form and the TSE-related scrapie form should follow the same pathway for all four species. In the preceding section, the much-discussed species barrier for infectious transmission of prion diseases (Prusiner, 1998) was placed in context with subtle local structure variations in this preserved scaffold. However, as present knowledge of structure–function correlations in prion proteins consists primarily of the observation that the same polypeptide chain can be found either in PrPC or PrPSc, and neither typical protein functions nor possible functional sites have been unambiguosly identified for PrPC, searches for physiologically relevant structural features have so far been highly speculative. Besides the considerations on the species barrier in Section III, examples of such searches include investigations of a possible structural basis of familial human TSEs (Liemann and Glockshuber, 1999; Riek et al., 1998) and the identification of epitopes for immune reactions in PrPC (Korth et al., 1997). Overall, however, because of the unusual circumstances of the discovery of PrPC, the lack of knowledge on “healthy” physiological functions of PrPC, and the remaining uncertainty about the exact role of the prion protein in TSE pathology, it appears that most or all physiologically relevant structure–function correlations of PrPC have yet to be identified.

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Besides the exciting prospect of future mining of the PrPC structures, which could be further supported by structure determinations of complexes with PrPC, there is also the outlook to structural biology with PrP forms that might be representative of some or all aspects of PrPSc. Considering the failure so far of obtaining suitable crystals for diffraction studies, NMR might again be the method of choice. For example, novel solution NMR techniques for large particles (Pervushin et al., 1997; Riek et al., 1999) might be applied with samples obtained through in vitro preparation of isotope-labeled aggregated forms of PrP along the lines that were successful for yeast prion-propagating proteins (King et al., 1997). Alternatively, solid state NMR techniques might open avenues for studies of aggregated forms of PrP (Heller et al., 1996; Antzutkin et al., 2000). ACKNOWLEDGMENTS We thank Mrs. M. Geier for the careful processing of the manuscript, and Dr. L. Calzolai, Dr. P. Güntert, T. Lührs, D. A. Lysek, C. von Schroetter, and Dr. R. Zahn for intellectual stimulation in our prion protein group. Our research projects in prion protein structural biology are supported by the ETH Zürich, the Schweizerischer Nationalfonds (projects 2-77-086.91 and 2-77560-97) and the BIOMED program of the European Commission (BBW-Nr. 97.0578-2).

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FOLDING DYNAMICS AND ENERGETICS OF RECOMBINANT PRION PROTEINS BY RUDI GLOCKSHUBER Institut für Molekularbiologie und Biophysik, Eidgenössische Technische Hochschule, Hönggerberg, CH-8093 Zürich, Switzerland

I. II. III. IV.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Folding of Recombinant PrPC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Role of the Single Disulfide Bond of PrP . . . . . . . . . . . . . . . . . . . . . . . . . . Influence of Point Mutations Linked with Inherited Human Prion Diseases on the Thermodynamic Stability of Recombinant PrPC . . . . . . . . . . . V. Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. INTRODUCTION Prions, the causative agents of transmissible spongiform encephalopathies (TSEs), such as Creutzfeldt-Jakob disease (CJD) in humans, scrapie in sheep, and bovine spongiform encephalopathy (BSE) in cattle, are essentially composed of PrPSc, the abnormal, oligomeric form of the host-encoded, monomeric, cellular prion protein PrPC. The protein-only hypothesis states that the prion is identical to PrPSc, and that PrPSc propagates through recruitment of PrPC from newly infected cells (Alper et al., 1967; Griffith, 1967; Prusiner, 1982; Prusiner, 1997). Numerous results support the protein-only hypothesis, which include the apparent absence of nucleic acids in prions, the dependence of prion susceptibility on the presence of PrP in the host (Büeler et al., 1993), and the association of point mutations in the human prion protein gene with inherited human TSEs (for reviews see Weissmann et al., 1996; Prusiner, 1997; Prusiner et al., 1998; Weissmann, 1999). Nevertheless, the experiment that would be considered by many scientists as the final proof of the protein-only hypothesis, i.e., the generation of prions from natural or recombinant PrPC in vitro, has not been reported so far (Horiuchi and Caughey, 1999). This brings structural and biophysical studies on PrPC and PrPSc into the center of interest, as they are expected to provide the basis for understanding the molecular mechanism of the conversion of PrPC into PrPSc. Mammalian prion proteins are secretory cell surface proteins of approximately 210 amino acids. All known mammalian PrP sequences are strikingly similar and pairs of sequences are generally more than 83 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

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90% identical (Schätzl et al., 1995; Wopfner et al., 1999). Futher characteristic features of mammalian PrPs are their posttranslational modifications, which include two N-glycosylation sites at Asn181 and Asn196, a single disulfide bond between Cys179 and Cys214, and a glycosylphosphatidylinositol (GPI) membrane anchor at the C-terminal residue 231 [residue 23 is the N-terminal amino acid of mature PrP; amino acid numbering according to human PrP (Schätzl et al., 1995)]. Although PrPC and PrPSc possess identical covalent structures (Hope et al., 1986; Stahl and Prusiner, 1991; Stahl et al., 1993), they differ considerably in their biochemical and biophysical properties. PrPC is monomeric, soluble in nondenaturing detergents, and sensitive to proteases, whereas PrPSc forms detergent-insoluble amyloid aggregates that are partially proteinase K-resistant. That the subunits of PrPSc are uniformly degraded to an N-terminally truncated form termed PrP(27–30) that retains infectivity and ranges from about residue 90 to residue 231 provides strong evidence that PrPSc is an ordered oligomer (Weissmann et al., 1996; Collinge et al., 1996; Raymond et al., 1997). Circular dichroism (CD) and infrared spectroscopy data showed that PrPC is rich in α-helical structure (Baldwin et al., 1994; Pan et al., 1993; Pergami et al., 1996), whereas PrPSc has a high β-sheet content (Caughey and Raymond, 1991; Gasset et al., 1993; Pan et al., 1993; Safar et al., 1993). Within the framework of the protein-only hypothesis, all these data suggest that mammalian prion diseases are caused by misfolding of PrPC and subsequent oligomerization to PrPSc. Several theoretical models for self-replication of PrPSc are presently discussed. The template assistance model (Prusiner, 1991) proposes an autocatalytic mechanism of PrPSc propagation via PrPC-PrPSc heterodimers, whereas the nucleation-polymerization model (Jarrett and Lansbury, 1993) postulates that formation of a PrPSc nucleus of critical size from small equilibrium quantities of PrPSc monomers is the critical event for PrPSc propagation. It follows that studies on the structure of PrPC and alternative conformations of the protein are crucial for understanding the molecular events underlying the formation of PrPSc. However, difficulties in purification of large amounts of PrPC from its natural source and the insolubility of PrPSc have long prevented high resolution structure analysis for both isoforms occurring in vivo. In 1996 and 1997, methods became available to produce large quantities of structurally homogeneous, monomeric recombinant PrP and PrP fragments in Escherichia coli either by secretory expression in the bacterial periplasm (Hornemann and Glockshuber, 1996) or by oxidative refolding from cytoplasmic incusion bodies (Mehlhorn et al., 1996; Donne et al., 1997; Hornemann et al., 1997; James et al., 1997; Zahn et

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al., 1997). Since then, the three-dimensional structures of the recombinant prion proteins from mouse (Riek et al., 1996; Riek et al., 1997), hamster (James et al., 1997; Donne et al., 1997), and humans (Zahn et al., 2000) have been determined in solution by nuclear magnetic resonance (NMR) spectroscopy. As expected from the high degree of sequence identity, all three structures proved to be very similar. Mammalian PrP is composed of two structurally distinct moieties. The C-terminal residues 125–231 form a globular domain with a unique fold consisting of three α helices and a short antiparallel β sheet, whereas the N-terminal segment 23–124 is flexibly disordered (Figs. 1 and 2, see color insert). The three-dimensional structures of the recombinant prion proteins are in full agreement with all previous physical data on the secondary structure of mammalian PrPC, and it is now generally accepted that the solution structures of the recombinant proteins, which lack all posttranslational modifications of mammalian PrPC except for the disulfide bond Cys179–Cys214, correspond to the structure of PrPC. Besides many structural insights into the effects of point mutations in human PrP that are linked with inherited TSEs (Riek et al., 1998) and the species barrier phenomenon in mammalian prion diseases (Billeter et al., 1997), the most important result of the structure determination is that the segment 90–124, which is flexible in PrPC, most likey adopts a defined structure in PrPSc subunits. This occurs because the segment 90–231 in PrPSc is resistant to proteolysis, whereas PrPC is rapidly degraded by proteinase K. This review focuses on recent biophysical studies on folding, stability, and alternative conformations of recombinant, mammalian prion proteins produced in E. coli. These include the fragments PrP(90–231) from hamster and human PrP, which represent the protease-resistant core of PrPSc, the full-length murine prion protein PrP(23–231), and the structured C-terminal domain PrP(121–231) of murine PrPC. This domain was initially identified by its resistance against proteolytic degradation during periplasmic expression of longer PrP fragments in the periplasm of E. coli (Hornemann and Glockshuber, 1996) and was the first PrP segment from which a three-dimensional structure could be obtained (Riek et al., 1996). II. FOLDING OF RECOMBINANT PrPC In this section, the folding and the presently known conformational states of disufide-intact recombinant prion proteins are discussed. Section III deals with biophysical studies on folding and aggregation of reduced

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Color Plates Legends FIG. 1. Ribbon representation of the three-dimensional structure of recombinant murine PrP(23–231) (Riek et al., 1997). The N-terminal segment of the protein (residues 23–123) is flexibly disordered. The C-terminal segment (residues 124–231) forms a unique tertiary structure consisting of three α helices (residues 144–154, 175–193, and 177–219), a short segment of helix-like stucture (residues 222–226) (all depicted in yellow), and a two-stranded antiparallel β sheet (residues 128–131 and 161–164, depicted in bluish-green). The single disulfide bond connecting Cys179 (helix 2) and Cys214 (helix 3) is shown in gray. The segment comprising residues 121–231 proved to be resistant against proteolytic degradation during functional expression in the periplasm of Escherichia coli and is termed PrP(121–231). The unstructured segment 23–120 is indicated by the black lines that represent connections between two neighboring Cα-atoms. The figure was generated with MOLMOL (Koradi et al., 1996). FIG. 2. Ribbon diagram of the refined NMR structure of murine PrP(121–231) (Riek et al., 1998); same color code for regular secondary structures as in Fig. 1), showing the side chains of residues that are exchanged in inherited human TSEs. Residues that are replaced in inherited Creutzfeldt-Jakob diseases (CJDs) are indicated in blue, and residues that were found to be replaced in patients with Gerstmann-SträusslerScheincker syndrome (GSS) are shown in red. The human polymorphism at residue 129 (Met or Val), depicted in gray, determines the phenotype of familial TSE associated with the mutation Asp178Asn (pink), which is either fatal familial insomnia (FFI) (Met129/Asn178) or inherited CJD (Val129/Asn178).

forms of recombinant PrP. On the basis of the known three-dimensional stuctures of mammalian prion proteins in solution, one would expect that folding of disulfide-intact (oxidized), full-length PrP(23–231) and its Nterminally truncated fragment PrP(90–231) is restricted to the structured C-terminal segment 125–231. Indeed, folding studies on the disulfideintact proteins murine PrP(23–231), murine PrP(121–231), and human PrP(90–231) yielded very similar results, in agreement with the working model that folding of the structured C-terminal domain is essentially independent of the flexibly disorded N-terminal tail. Table I summarizes the present data on the folding of oxidized, recombinant prion proteins. In the range of pH 4.0 and 8.0 and in the presence of low ionic strength at pH 4 to 5, guanidinium chloride (GdmCl) and urea-induced equilibrium unfolding experiments revealed that murine PrP(121–231), human PrP(90–231), and murine PrP(23–231) show completely reversible transitions that are consistent with the two-state model of folding (i.e., a mixture of completely unfolded and completely folded molecules at any given denaturant concentration) (Hornemann and Glockshuber, 1996; Swietnicki et al., 1997; Swietnicki et al., 1998; Hosszu et al., 1999; Liemann and Glockshuber, 1999; Jackson et al., 1999; Cereghetti et al., 2000) (cf. Fig. 4A). A comparison between the urea-induced unfolding transitions of full-length murine PrP(23–231) and its isolated C-terminal domain PrP(121–231) showed that the full-length protein was

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TABLE I Properties of Various Conformational States of Recombinant Murine PrP(23–231) and PrP(121–231), and Comparison with Other Studies on the Fragments 90–231 from Syrian Hamster and Human PrP Protein Oxidized mPrP mPrP(121–231)a and mPrP(23–231)b mPrP(121–231)c and mPrP(23–231)d mPrP(23–231) Reduced mPrP mPrP(23–231)e mPrP(23–231)e mPrP(23–231)e

Conditions

Properties

pH 4–8 (low ionic strength at pH 4–5) pH 4.0, 3.5 M urea, ionic strength > 100 mM, protein concentration > 20 µM pH 7.0, ionic strength > 0.3 M

monomeric, α-helical, two-state folding transitionsf acid-induced unfolding intermediate, β-sheet-like CD spectra, oligomericg unspecific aggregation

pH 4.0–6.0, low ionic strength

β-sheetlike; pH-dependent structural plasticityh amyloid-like aggregatesi α-helical,j but less structured than oxidized mPrP(23–231), unspecific aggregation below 3 M ureae

pH 4.0, ionic strength > 0.2 M pH 7.4, 3.0 M urea

Hornemann and Glockshuber, 1996. Liemann and Glockshuber, 1999. c Hornemann and Glockshuber, 1998. d Cereghetti et al., 2000. e Zobeley et al., 2000. f Two-state transitions under similar conditions were also observed for human PrP(91–231) (Jackson et al., 1999b; Hosszu et al., 1999) and human PrP(90–231) (Swietnicki et al., 1997; Swietnicki et al., 1998). g Analogous intermediates have been observed for syrian hamster PrP(90–231) in 2–2.5 M GdmCl at pH 5.0 (Zhang et al., 1997), human PrP(91–231) in 1–1.5 M GdmCl at pH 4.0 and protein concentrations of 62 µM (Jackson et al., 1999a, 1999b) and human PrP(90–231) (about 6 µM) in 1–1.5 M GdmCl at pH 3.6 (Swietnicki et al., 1997; Swietnicki et al., 1998). Prolonged incubation of oxidized human PrP(90–231) in 1 M GdmCl between pH 3.6 and 5.0 yielded both amorphous and fibrillar aggregates rich in β-sheet structure that showed increased proteinase K resistance (Swietnicki et al., 2000). In addition, β-sheetlike CD spectra were also abtained after thermal unfolding and cooling of oxidized hamster PrP(90–231) (Zhang et al., 1997) and human PrP(91–231) (Jackson et al., 1999b). h β-sheet conformations at acidic pH, caused by reduction of the disulfide bond, were first reported for reduced hamster PrP(90–231) (Mehlhorn et al., 1996) and reduced human PrP(91–231) (Jackson et al., 1999a, 1999b). i Aggregation of reduced PrP to amyloid at pH 4.0 and ionic strengths > 0.1 M was first reported for human PrP(91–231) (Jackson et al., 1999a) j An α-helical CD spectrum practically identical to that of the oxidized protein was reported for reduced human PrP(91–231) after refolding at pH 8.0 (Jackson et al., 1999b). a b

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FIG. 3. Far–UV circular dichroism (CD) spectra of different conformational states of recombinant murine PrP(23–231) and PrP(121–231) that can be populated in solution: (1) oxidized PrP(121–231) at 4.0, (2) oxidized PrP(23–231) at pH 7.0, (3) acid-induced unfolding intermediate of oxidized PrP(121–231) in 3.5 M urea at pH. 4.0 (ionic strength: 88 mM; protein concentration: 30 µM), (4) reduced PrP(23–231) at pH 4.0 and 10 mM ionic strength, and (5) reduced PrP(23–231) in 3 M urea at pH 7.4.

FIG. 4. Urea-induced equilibrium unfolding transitions of murine PrP(121–231) at 22°C. (A) Dependence of reversible, urea-induced unfolding on pH. Open symbols represent refolding experiments, and closed symbols represent unfolding experiments. (B) Dependence on ionic strength of the formation of the acid-induced unfolding intermediate of PrP(121–231) at pH 4.0 and a protein concentration of 29 µM. Unfolding experiments were performed at 22°C in 50 mM formic acid/NaOH, pH 4.0 (ionic strength: 32 mM), containing 0 M (❍), 50 mM (❑), 100 mM (∆), or 150 mM (■) sodium chloride.

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reproducibly slightly less stable (∆Gfold = –25.5 kJ mol–1 compared with –29.7 kJ mol–1) (Liemann and Glockshuber, 1999). Both proteins have identical midpoints of tansitions (6.2–6.3 M urea), but the cooperativity of folding is slightly lower in the case of the full-length protein. As no interactions between the C-terminal domain and the unstructured N-terminal segment were observed in PrP(23–231) (Riek et al., 1997), the lower stability of the full-length protein may result from contacts between residues from the N- and C-terminal parts of PrP(23–231) in the unfolded state. Such a residual structure in the unfolded state would reduce the difference in the solvent-accessible surface area between the folded and unfolded state and decrease folding cooperativity (Myers et al., 1995). The complete reversibility of unfolding of PrP(121–231), PrP(90–231) and PrP(23–231) has principal implications within the framework of the protein only hypothesis if one extrapolates the results from the recombiant proteins to natural PrPC. If PrPC and PrPSc indeed have identical covalent stuctures (Stahl and Prusiner, 1991), both PrPC and PrPSc will yield identical unfolded forms in the presence of high concentrations of the denaturants urea and GdmCl. This predicts that, after reconstitution in vitro, one would always obtain folded PrPC, independent of whether the experiment was started from PrPC or PrPSc. This would explain why all attempts to reconstitute infectivity after complete solubilization of infectious PrPSc with high concentrations of GdmCl or urea have failed (Prusiner et al., 1993). Further studies fully confirmed the two-state character of folding of oxidized recombinant PrP. Analysis of the kinetics of folding of the tryptophan variant F175W of murine PrP(121–231) at pH 7.0 revealed extremely dynamic unfolding equilibria, which could be observed only with stopped-flow fluorescence at 4°C (Wildegger et al., 1999). The kinetics of unfolding and refolding of this mutant domain were in complete agreement with the two-state model and the equilibrium unfolding data. No kinetic intermediates were observed during refolding of urea-denatured PrP(121–231)-F175W. Indeed, folding of the structured C-terminal PrP domain is one of the fastest protein folding reactions described so far, with an extrapolated half-life of folding of 170 µs (half-life of unfolding at pH 7.0 and 4°C: 4.6 min) (Wildegger et al., 1999) (Fig. 5). This almost certainly excludes the recruitment of transiently populated PrP folding intermediates as precursors of PrPSc, which would be expected to cause significantly slower folding. Kinetic folding intermediates have been proposed for amyloid formation of human lysozyme mutants (Booth et al., 1997). The two-state character of folding of recombinant PrP implies that the unfolded state of PrPC could be the starting point of a structural

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FIG. 5. Kinetics of folding of murine PrP(121–231) (variant F175W) at pH 7.0 and 4°C, measured by stopped-flow fluorescence. PrP(121–231)-F175W was unfolded with 8 M urea and diluted with refolding buffer to a final concentration of 3.7 M urea. The extrapolated value at t = 0 s corresponds to the expected value from the urea-induced equilibrium transition (excitation: 280 nm; emission: > 320 nm).

transition to PrPSc under physiological conditions. The same result was obtained from the analysis of backbone amide hydrogen exchange in human PrP(91–231) by NMR spectroscopy (Hosszu et al., 1999). These studies fully confirmed the two-state character of PrP folding at pH 5.5 and revealed that only a small portion of residual structure within about 10 residues around the disulfide bond is retained in the unfolded state (Hosszu et al., 1999). Overall, the C-terminal domain of PrPC is a comparably rather stable protein, shows perfect two-state behavior of folding at low ionic strength, and is as “normal and well-behaved” as a protein can be. No property of the domain at physiological pH indicates that it is capable of adopting an entirely different conformation. Such an alternative conformation was first observed for a recombinant prion protein as a plateau phase in the GdmCl-induced unfolding transition of hamster PrP(90–231) at pH 5.0 (Zang et al., 1997). Studies on the pH-dependence of PrP folding then showed that human PrP(90–231) at pH 3.6–5.0 in the presence of GdmCl (Swietnicki et al., 1997; Jackson et al., 1999b) and murine PrP(121–231) and PrP(23–231) at pH 4.0–4.5 in the presence of urea (Hornemann and Glockshuber, 1998; Cereghetti et al., 2000) (Fig. 3,

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trace 3; Fig. 4A) populate an acid-induced unfolding intermediate at medium denaturant concentration. All these intermediates showed a strong content of β-sheet structure in far-UV CD measurements. In the case of murine PrP(121–231), the apparent pKa of the transition from two-state to three-state folding characteristics in the presence of urea is 4.5, indicating that protonation of acidic side chains is responsible for stabilization of the intermediate (Hornemann and Glockshuber, 1998). Further studies on the formation of the scrapie-like, acid-induced unfolding intermediate in the presence of urea, using murine PrP(121–231) and full-length PrP(23–231) as a model, revealed the following (Cereghetti et al., 2000): (1) Formation of the intermediate is an intrinsic property of the C-terminal PrP domain 121–231 and independent of the flexible Nterminal segment 23–120, (2) the intermediate is an oligomer and only significantly populated at protein concentrations > 20 µM at pH 4.0, and (3) population of the intermediate requires an ionic strength > 75 to 100 mM (Fig. 4B). Salt stabilization of the intermediate may occur either by hydrophobic interactions within the intermediate, or by weakening of repulsive interactions between subunits in the intermediate. In the case of human PrP(91–231), the apparent molecular mass of a dimer was obtained for the intermediate when populated in 1 M GdmCl at pH 4.0 (Jackson et al., 1999a). Prolonged incubation of the acid-induced unfolding intermediate of oxidized human PrP(90–231) in 1 M GdmCl in the range of pH 3.6–5.0 yieled specific fibrillar aggregates that showed increased proteinase K resistance (Swietnicki et al., 2000). Whether these aggregates are infectious is not yet known, but they show essential characteristics of PrPSc. As PrPSc accumulates in endosomes of scrapie-infected cells where acidic pH values of 4.0 to 6.0 are prevalent (Lee et al., 1996), it is tempting to speculate that a change from physiological to acidic pH during endocytosis of PrPC triggers its conversion to PrPSc (Hornemann and Glockshuber, 1998; Kelly, 1998a). In summary, folding of oxidized, recombinant PrP can be described by Scheme 1. pH 4–8, low ionic strength at pH 4–5 pH 4, Ionic strength > 50 mM [PrP] > 20 µM

N nN

U In

nU

Scheme 1: Dependence of folding of murine PrP(121–231) and PrP(23–231) on pH and ionic strength. N, I, and U correspond to the native, intermediate and unfolded state, respectively. Data on human PrP(91–231) indicate that the acid-induced equilibrium intermediate In (populated in the presence of GdmCl) is dimeric (n = 2) (Jackson et al., 1999a).

Finally, it was found for oxidized human PrP(91–231) that the protein could be converted from an α-helical to a β-sheet-rich conformation after

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thermal unfolding and cooling (Jackson et al., 1999b). Whether this heatinduced β-sheet conformation has similarities with the acid-induced unfolding intermediate of recombinant PrP or the reduced state of PrP at low pH and low ionic strength (see later) still has to be established. III. THE ROLE OF THE SINGLE DISULFIDE BOND OF PrP The hypothesis that mammalian prion diseases are caused only by an alternative conformation of the prion protein is essentially based on the fact that no differences between the covalent structures of PrPC and PrPSc could be detected so far (Stahl and Prusiner, 1991). PrPSc subunits thus bear all posttranslational modifications that are found in PrPC, even though the fractions of the unglycosylated, monoglycosylated and doubly glycosylated forms of PrP differ in a prion strain-specific manner (Bessen et al., 1995; Collinge et al., 1996). The single intramolecular disulfide bond Cys179–Cys214 of PrP is also assumed to be quantitatively formed in both PrPC and PrPSc, as denaturant-solubilized PrPSc only shows reactivity with thiol-specific reagents after treatment with reducing agents (Turk et al., 1988). As mentioned previously, disulfide bond formation in vivo or oxidative refolding of PrP in vitro has been essential for producing recombinant PrPC for structure determination. In addition, the oxidized state of PrP was required for the PrPSc-mediated formation of protease-resistant PrP from PrPC in vitro (Herrmann and Caughey, 1998), and incubation of mouse prions with 2-mercaptoethanol in the presence of SDS decreased scrapie infectivity (Somerville et al., 1980). All these data suggest that the disulfide bond is permanently present during the conversion of PrPC into PrPSc. However, this does not nessecarily have to be the case because the disulfide bond in PrP may become reduced transiently during the conversion into PrPSc, and reoxidation of the cysteines may occur at a later stage of PrPSc formation (e.g., at the level of PrPSc after oligomerization of PrPSc subunits). This idea was first presented by Mehlhorn et al. (1996) who observed that reduced, recombinant hamster PrP(90–231), in contrast to the oxidized protein, showed β-sheetlike CD spectra. Further studies revealed that reduced hamster PrP(90–231) tends to oligomerize and aggregate at pH 6.5 (Zhang et al., 1997). More recently, the pH-dependence of the physical properties of reduced human PrP(91–231) was investigated in great detail (Jackson et al., 1999a). Although oxidized human PrP(91–231) shows the expected α-helical characteristics over a wide pH range, the reduced

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protein can exist in at least two different states. At low pH (pH 4.0) and low ionic strength, reduced PrP(91–231) is soluble, shows β-sheet-like far-UV CD spectra, possesses a significant degree of tertiary structure (as shown by a characteristic near-UV CD signal), and shows the apparent molecular mass of the monomer in gel filtration experiments. However, the tertiary structure content of the reduced protein is significantly lower compared with oxidized PrP(91–231) and reminiscent of a molten globule state, as evidenced by a strong loss of chemical shift dispersion in 1H-15N heteronuclear single-quantum coherence (HSQC) NMR spectra (Jackson et al., 1999a). Increasing the ionic strength to > 100 mM at pH 4.0 triggers specific aggregation of reduced PrP(91–231), eventually leading to amyloid-like fibers showing limited proteinase K resistance relative to the oxidized protein (Jackson et al., 1999a). Of interest, monomeric, reduced PrP(91–231) at low ionic strength already shows protease resistance to almost the same extent (Jackson et al., 1999a). Overall, the protease resistance of fibers of reduced PrP(91–231) is significantly lower than that of PrPSc. Nevertheless, these fibers, produced in vitro from recombinant protein, exhibit many similarities with PrPSc. Whether these aggregates are infectious or become infectious after reoxidation still remains to be established. In vitro results similar to those on reduced human PrP(91–231) described previously were obtained for the reduced full-length murine prion protein PrP(23–231), which also shows β-sheetlike spectroscopic properties at acidic pH (cf. Fig. 3) and aggregates to fibrils under practically the same conditions as human PrP(91–231) (Zobeley et al. 2000). Reduced PrP(23–231) also populates another, new conformation in solution at pH 7.4, which differs significantly in its far-UV CD spectra from the oxidized protein and the reduced protein at pH 4.0 (Fig. 3, trace 5). Surprinsingly, the reduced protein at pH 7.4 proved to be almost as stable as the oxidized protein and showed cooperative, onestep unfolding transitions in the presence of urea with similar cooperativities (–23.7 kJ mol–1 compared with –26.7 kJ mol–1, respectively). However, reduced PrP(23–231) has a strong tendency to aggregate at pH 7.4 in the absence of urea. This aggregation process is unspecific and does not lead to the fibrils observed at pH 4.0 (Zobeley et al., 2000). In the known NMR structures of murine, hamster and human recombinant PrPC, the invariant disulfide bond Cys179–Cys214 is entirely buried in the interior of the structured, C-terminal domain and not accessible to external reductants. In accordance with the structural data is the finding that the disulfide bond of murine PrP(23–231) is extremely resistant against the strong reductant dithiothreitol (DTT).

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The half-time of reduction by 50 mM DTT in the range of pH 4.0–9.0 and 37°C is > 10 h, indicating that unfolding of PrP must precede disulfide bond reduction (Zobeley et al., 2000). This raises the question of how PrPC can become reduced in vivo, in particular in the acidic environment of endosomes where disulfide exchange reactions are extremely slow and PrPSc appears to accumulate. One possibility might be endocytic enzymes such as the recenty described γ-interferoninducible lysosomal thiol reductase (GILT), which is optimally active as catalyst of protein disulfide bond reduction at pH 4 to 5 (Arunachalam et al., 2000). In accordance with this hypothesis is the finding that disulfide oxidoreductases such as DsbA from E. coli efficiently catalyzed reduction of PrP(23–231) by DTT both at physiological and acidic pH (Zobeley et al., 2000). Almost quantitative reoxidation of aggregates of reduced PrP(23–231) by the dithiol oxidant diamide could be achieved. However, in contast to the amyloids of reduced human PrP(91–231), an increased protease resistance was not observed for the reduced and the reoxidized amyloids of murine PrP(23–231) (Zobeley et al., 2000). In summary, the recombinant proteins murine PrP(23–231), human PrP(91–231), and hamster PrP(90–231) all share an unusual feature that has so far not been observed in other proteins (Jackson et al., 1999a): Their oxidized and reduced forms adopt entirely different, stable tertiary structures that fold cooperatively. The structural features of the oxidized proteins correspond to those of PrPC, and the properties of the reduced forms are reminiscent of PrPSc. This provokes speculations about a tansient reduction of the disulfide bond of PrP during the conversion process in vivo. A role of reduced PrP in PrPSc formation is particuarly supported by the recent finding that high-level expression of unglycosylated murine PrP(23–231) in the reducing environment of the yeast cytoplasm yields PrP aggregates that are insoluble in the detergent Sarkosyl and show a proteinase K resistance pattern similar to that of PrPSc (degradation of the N-terminal residues 23 to ~90; protease resistant core: ~90–231) (Ma and Lindquist, 1999). The authors suggest that retrograde transport to the cytoplasm of misfolded PrP in the endoplasmic reticulum or transmembrane forms of PrP (Hedge et al., 1998), followed by aggregation of reduced PrP, might be a possible initial event of PrPSc nucleus formation (Ma and Lindquist, 1999). Aggregation to an ordered, protease-resistant oligomer has not been reported so far for aggregates of reduced PrP produced in the E. coli cytoplasm. Additional cellular factors present only in eukaryotic cells might therefore be required for formation of PrPSc-like aggregates of reduced, recombinant PrP. A small oligomer of reduced PrP serving as

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a nucleus for growth of large, oxidized PrPSc oligomers (Jackson et al., 1999a, Ma and Lindquist, 1999) would be an explanation for the apparent absence of free thiol groups in PrPSc, because the fraction of reduced PrP in the oxidizd PrPSc oligomer may simply be too small for being detectable. It is obvious that the most important experiment for clarifying the possible role of reduced PrP in PrPSc formation is testing reduced PrP aggregates that were generated in vitro or in vivo in heterologous expression systems for infectivity. IV. INFLUENCE OF POINT MUTATIONS LINKED WITH INHERITED HUMAN PRION DISEASES ON THE THERMODYNAMIC STABILITY OF RECOMBINANT PrPC The fact that all forms of inherited human prion diseases known so far are linked with dominant mutations in the gene encoding human PrP has been used as a strong argument in favor of the protein-only hypothesis, as the infectious prion agent spontaneously develops in affected individuals (for reviews, see Prusiner, 1997; Prusiner et al., 1998). Three different phenotypes of inherited human prion dieseases are known: the Gerstmann-Sträussler-Scheincker syndrome (GSS), familial Creutzfeldt-Jakob disease (CJD), and fatal familial insomnia (FFI). Besides point mutations in the human PrP gene, there are also insertions of additional octapeptide repeats that cause inherited TSEs in humans (Fig. 6). As the octapeptide repeat segments, the supposed Cu2+ binding regions of PrP (Viles et al., 1999), are not required for generation and propagation of prions (Fischer et al., 1996) and not contained in the infectious, protease-resistant core of PrPSc, the mechanism by which additional octapeptides lead to spontaneous prion formation is not obvious. In contrast, all single amino acid replacements that cause inherited human TSEs are located within the minimum infectious unit of PrPSc (i.e., within segment 90–231) (Fig. 6). Different mechanisms for the spontanous generation of prions in familial human TSEs through point mutations in PrP have been postulated (Cohen, 1999). A mechanism that has been discussed for many years proposes that the single amino acid replacements within mature PrP either decrease the thermodynamic stability of PrPC and thereby facilitate its conversion to PrPSc (Cohen et al., 1994; Huang et al., 1995) and/or increase the stability of PrPSc (Cohen, 1999). A second model suggests that the mutations accelerate the formation PrPSc by decreasing the activation energy barrier for the conformational transition (Cohen, 1999) or accelerate the assembly of PrPSc monomers (Liemann and Glockshuber, 1999). As PrPSc formation is favored at high concentrations of PrPC (Büeler et al., 1993), it could also be that some amino acid

FIG. 6. Influence of amino acid replacements associated with inherited human TSEs on the thermodynamic stability of recombinant murine PrP(121–231) at pH 7.0 and 22°C (Liemann and Glockshuber, 1999). The upper panel indicates the amino acid replacements in mature human PrP that are linked with inherited prion diseases in humans. The lower panel shows the difference between the free energy of unfolding of wild-type murine PrP(121–231) and the corresponding variant (∆∆G). The difference is defined such that a destabilization relative to the wild-type leads to positive values of ∆∆G. The free energy of folding of wild-type murine PrP(121–231) at pH 7.0 and 22°C is –29.7 ± 1.0 kJ mol–1. The maximum error of the measured ∆∆G values is ± 2.6 kJ mol–1. Variants of PrP(121–231) that could not be expressed in a soluble form in the E. coli periplasm and formed periplasmic inclusion bodies are marked by asterisks. ∆∆G for the replacement E200K has also been determined for human PrP(90–231) with guanidinium chloride as denaturant, and has a value of 4.1 ± 2.6 kJ mol–1 (Swietnicki et al., 1998, dashed bar frame).

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replacements affect the intracellular turnover of PrP through a variety of mechanisms. Finally, the amino acid replacements might also improve the interaction between PrP and the postulated host factor “protein X,” which is supposed to promote prion propagation in a species-specific manner (Telling et al., 1995; Kaneko et al., 1997). Both the thermodynamic and kinetic models of facilitated, spontaneous prion propagation in inherited TSEs are in accordance with the observation that sporadic CJD, which does not segregate with mutations in the prion protein gene, has a very late onset (> 65 years), whereas all inherited TSEs are characterized by a comparably early onset at the age of 30 to 40 years. However, mapping of the TSE-related amino acid replacements in human PrP onto the solution structure of recombinant PrP immediately suggests that destabilization of PrPC is unlikely to be the general mechanism of inherited prion generation because the three GSS-related mutations—P102L, P105L, and A117V—are located in the flexibly disordered segment of PrPC, and mutations in the unstructured polypeptide segment are not expected to influence the total stability of PrPC. It follows that the thermodynamic hypothesis of PrPC destabilization can apply only to the structured domain PrP(121–231). Indeed, the thermodynamic stability of the variant P102L of human PrP(90–231) was indistinguishable within experimental error from that of the wild-type protein (Swietnicki et al., 1998). Figures 2 and 6 show that the eight disease-related replacements within the C-terminal domain of PrPC are located in the segment containing α helices 2 and 3, which form the scaffold of the structured domain (Riek et al., 1996; Riek et al., 1997). A detailed structural analysis of the C-terminal domain of PrP indicated that only four of eight amino acid replacements in PrP(121–231) are likey to be destabilizing, namely D178N (loss of a salt bridge between D178 and R164, influence on the charged hydrogen bond between the carboxylate of D178 and the side chain hydroxyl group of Y128), T183A (loss of two hydrogen bonds between the hydroxyl group of T183 and the main chain carbonyl group of C179 and the main chain amide group of Y162), Q217R (loss of a hydrogen bond between the side chain amide of Q217 and the main chain amide of A133), and the replacement F198S (creation of a relatively large cavity) (Riek et al., 1998). To test the influence of the eight TSE causing point mutations on the stability of the structured C-terminal domain of human PrP, murine PrP(121–231) was used as a model. In addition to its high strucutral similarity and 94% sequence identity to human PrP(121–231), all eight residues whose replacements segregate with inherited TSEs are identical in wild-type human and murine PrP, and the side chains that form

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direct contacts with these residues in the structures of murine and human PrP(121–231) are also identical (Riek et al., 1998; Zahn et al., 1999). Murine PrP(121–231) thus represents a reasonable model system to study the effect of amino acid replacements in the structured domain of human PrPC. The eight amino acid replacements were introduced individually into murine PrP(121–231), and all variants showed the same far- and near-UV CD spectra as the wild-type domain, demonstrating that none of these mutations a priori induces a PrPSc-like conformation with increased β-sheet and decreased α-helix content (Liemann and Glockshuber, 1999). However, exactly those variants that had been predicted to be destabilized (D178N, T183A, F198S and Q217R) showed a strong tendency to aggregate during secretory expression in E. coli. They formed periplasmic inclusion bodies and had to be refolded before biophysical analysis (Fig. 6) (Liemann and Glockshuber, 1999). Treatment of the periplasmic inclusion bodies with proteinase K leads to complete degradation of the aggregates, showing that they had no characteristic similarities to PrPSc and were unspecific aggregates (G. Cereghetti and R. Glockshuber, unpublished results). Urea-induced equilibrium unfolding experiments proved that indeed only the replacements D178N, T183A, F198S, and Q217R were destabilizing, whereas the exchanges V180I, E220K, and V210I, and the human polymorphism replacement M129V essentially had no influence on the thermodynamic stability of PrP(121–231) (Fig. 6) (Liemann and Glockshuber, 1999). Like the wild-type domain, all eight variants of PrP(121–231) showed completely reversible one-step transitions at pH 7.0 that were consistent with the two-state model of folding. Destabilizing amino acid replacements essentially shifted the transitions midpoints to lower denaturant concentration, whereas the cooperativities of the tansistions were only affected to small extent. The replacement T183A proved to be most destabilizing, and the corresponding PrP(121–231) variant (∆Gfold = –10.4 kJ mol–1 was about three times less stable than wild-type PrP(121–231) (∆Gfold = –29.7 kJ mol–1) (Fig. 6) (Liemann and Glockshuber, 1999). Overall, like the mutations in the flexible tail of PrPC, the analysis of the variants of the structured PrPC domain showed that point mutations in the human PrP associated with inherited TSEs are not necessarily destabilizing. It could well be that the thermodynamic model of prion generation in inherited TSEs applies to the mutations D178N, T183A, F198S, and Q217R; but other mechanisms are likely to underlie prion generation in the case of the replacements V180I, E220K and V210I. Of importance, there is no correlation between the stabilities of the variants of PrP(121–231) and the disease phenotypes. This is particu-

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larly obvious in the case of the mutation D178N, which, in conjunction with the polymorphism at residue 129 (Met or Val), determines the phenotype of familial TSE associated with the D178N mutation (FFI in the case of M129/N178 or inherited CJD for V129/N178) (Goldfarb et al., 1992). The replacement M129V alone had no effect on protein stability, and the same was observed for the polymorphism replacement in the D178N variant (Fig. 6). Studies on the expression of human PrP genes harboring the D178N mutation (Petersen et al., 1996) and the Q217R mutation (Singh et al., 1997) in human neuroblastoma cells are in agreement with the thermodynamic stabilities of these corresponding mutant mouse PrP(121–231) fragments. Proteins with the Asp178Asn exchange were unstable in vivo (Petersen et al., 1996), and temperature-dependent protein misfolding was observed in the case of the Gln217Arg variant (Singh et al., 1997). For several murine PrP proteins carrying different mutations associated with inherited TSEs, the production in eukaryotic cells has been reported. Variants of murine PrP variants P102L, D178N/M129, T183A, and E200K expressed in Chinese hamster ovary (CHO) cells displayed a number of PrPSc-like properties such as detergent insolubility and protease resistance, and the amino acid exchange T183A blocked the delivery of the protein to the cell surface, probably owing to inactivation of the N-glycosylation site at Asn181 (Lehmann and Harris, 1995; Lehmann and Harris, 1996a; Lehmann and Harris, 1996b; Daude et al., 1997; Lehmann and Harris 1997). Although these mutant PrPs are distinguished from each other by several biochemical features such as the size of their protease-resistant cores or the glycosylation patterns (Lehmann and Harris, 1996a, b), the thermodynamic stabilities of the analogous variants of PrP(121–231) do not correlate with these properties. In summary, it is presently not possibe to deduce a clear-cut molecular mechanism of spontaneous prion generation in inherited TSEs from the thermodynamic stabilities of mutant PrP variants and data on their expression in cultured cells. Finally, an important feature of prions in familial TSEs may be the stoichiometry and subunit composition of PrPSc isolated from affected individuals. Analysis of the allelic origin of the protease-resistant PrPSc oligomer isolated from patients with inherited prion diseases yielded different results for individual mutations. In the case of the mutations at positions 102, 178, 198, 200, and 217, only the mutant PrP forms protease-resistant PrPSc (Kitamoto et al., 1991; Tagliavini et al., 1994; Barbanti et al., 1996; Gabizon et al., 1996; Chen et al., 1997). In contrast, the mutation at position 210 (Silvestrini et al., 1997) and also the insertion

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of five or six additional octapeptide repeats (Chen et al., 1997) led to heterooligomeric, protease-resistant PrPSc deposits in the brain of the patients. Consequently, there is no correlation between the thermodynamic stability of PrPC and the tendency of mutated PrPs to form PrPSc homooligomers or heterooligomers. Instead, the homooligomeric or heterooligomeric association state of PrPSc in inherited TSEs may be determined by the intermolecular interactions in a supposed heterodimeric PrPC/PrPSc intermediate (Prusiner, 1991) or by the abilities of wild-type and mutated PrPSc monomers to assemble into a proteaseresistant, regular array of PrPSc subunits (Jarrett and Lansbury, 1993). V. CONCLUSIONS Two different experiments have succeeded so far in de novo generating protease-resistant oligomers of recombinant mammalian PrP with PrPSc-like properties in vitro. These are the generation of fibrils of oxidized human PrP(90–231), starting from conditions where the β-sheetrich, acid-induced unfolding intermediate of PrP is stongly populated (Swietnicki et al., 2000), and the in vitro formation of fibrils of reduced human PrP(91–231) and reduced murine PrP(23–231) (Jackson et al., 1999a; Zobeley et al., 2000). However, no infectivity in preparations of these fibrils has yet been reported. In contrast to the well-established de novo generation of self-propagating aggregates of the yeast nonmendelian prion factors Sup35p (King et al., 1997; Glover et al., 1997; DePace et al., 1998; Santoso et al., 2000) and Ure2p (Taylor et al., 1999), an important feature of self-propagating aggregates, namely, the generation of new aggregates by inoculation with catalytic amounts of preexisiting aggregates, could so far not be performed successfully with recombinant mammalian PrP. This could be due to the fact that essential components for formation of infectious PrPSc aggregates have been lacking in these in vitro experiments, such as sphingolipids, which were found to be associated with PrPSc (Klein et al., 1998), and complex polysaccharides, which have been proposed to form the scaffold of PrPSc (Appel et al., 1999). Another important difference between the presently perfomed in vitro experiments in solution and the situation in vivo is that PrPSc is most likely formed at the surface of membranes, either within caveolaelike domains (Kaneko et al., 1997) or in endosomes (Borchelt et al., 1992; Arnold et al., 1995), whereas the recombinant proteins lack a Cterminal transmembrane anchor. It is also striking that among the numerous amyloid diseases in humans including Alzheimer’s disease,

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only PrPSc amyloids are infectious, and that PrP is simultaneously the only amyoidogenic protein with a membrane anchor (Kelly, 1998b). It could thus well be that incorporation of PrPC into membranes, which is expected to increase the effective concentrations of PrPSc precursers and to preorient aggregating subunits relative to each other, is required for in vitro formation of infectious PrPSc. In addition, recombinant human PrP(23–231) has been shown to interact with synthetic membrane vesicles and undergo slight conformational changes on membrane binding (Morillas et al., 1999). Membrane attachment may thus favor alternative conformations of PrPC and its transitions to PrPSc. Moreover, natural protein ligands or cofactors such as Cu2+ions, which are supposed to be natural ligands of PrPC in vivo (Brown et al., 1997), bind to the histidine-rich octapeptide repeat region (residues 51–91) in the unstructured region of PrPC (Viles et al., 1999; Whittal et al., 2000) and also influence PrPSc conformation (Wadsworth et al., 1999). Regarding the multitude of paramters that can be varied in in vitro conversion experiments, it may take time and good luck to generate prions de novo from natural PrPC or recombinant PrP. ACKNOWLEDGMENTS This work was supported by the Schweizerischer Nationalsfonds (Project 438+50285) and the Schweizerisches Bundesamt für Bildung und Wissenschaft (Project ‘97.0578-1). Special thanks to Eva Zobeley for discussions and careful reading of the manuscript.

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SIMULATIONS AND COMPUTATIONAL ANALYSES OF PRION PROTEIN CONFORMATIONS BY DARWIN O.V. ALONSO AND VALERIE DAGGETT Department of Medicinal Chemistry, University of Washington, Seattle, Washington 98195

I. PrP Conformational Transitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Predictions of PrP Structures and Studies of Peptide Fragments to Test the Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Prediction of the Structure of PrPC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Prediction of the Structure of PrPSc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Peptide Fragments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. PrPC Structural Models from NMR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Detailed Modeling Studies Based on the NMR Models . . . . . . . . . . . . . . . . . . A. Static Modeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Detailed Atomic-level Molecular Dynamics Simulations of PrP . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. PrP CONFORMATIONAL TRANSITIONS The folding of most globular proteins results in a structure with a stable conformation. Although proteins undergo conformational changes on ligand binding or posttranslational modification, the gross secondary structure framework generally remains intact. When globular proteins fold incorrectly as a result of an amino acid mutation, chemical modification, change in the environment, or other unknown factors, the proteins are generally degraded or they aggregate. In the limiting case of degradation, this may result in the absence of a functional protein. In the case of aggregation of misfolded forms of a protein, limited polymerization that impairs biological function may occur, as is the case with sickle cell anemia. Or, there may be more widespread aggregation leading to plaque formation, as seen in a variety of amyloid diseases. Given the nature of the conformational changes involved in these disease states, they can be considered “folding diseases” and they fall under the “toxic conformer” heading of Thomas et al. (1995). Characterization of the conformational properties of peptides and proteins at the atomic level is challenging when using experimental means and becomes particularly difficult when the process involves aggregation. For example, while one may be able to investigate the structure of a protein in solution under a particular set of conditions, changes in the conditions necessary to elicit a conformational change relevant to amyloidosis generally lead to aggregation and insolubility, 107 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

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such that different experimental techniques must be used to study the two endpoints. This is difficult enough, but it is nearly impossible to characterize the actual conformational transition or folding/unfolding process between these endpoints. Given that protein folding and misfolding are of such widespread importance to human health and the fact that experimental approaches provide only limited amounts of information on the structural transitions and interactions occurring during these processes, modeling and computer simulation methods are used to complement and extend experiment. Prions are transmissible pathogens that cause fatal neurodegenerative diseases of the central nervous system in humans and animals (Prusiner, 1991, 1996). In humans, prion diseases have been subdivided into four classes and are referred to as kuru, Creutzfeldt-Jakob disease (CJD), Gerstmann-Sträussler-Scheinker disease (GSS), and fatal familial insomnia (FFI) (DeArmond and Prusiner, 1995; Prusiner, 1998). These diseases, as discussed in more depth elsewhere in this volume, can be sporadic, inherited, or infectious (or iatrogenic) disorders. They differ from other infectious diseases in that the pathogen is a proteinaceous particle, termed a prion (Prusiner, 1991, 1996, 1998), and the essential component of prions is the scrapie prion protein (PrPSc). PrPSc is chemically indistinguishable from the normal, cellular prion protein (PrPC) (Stahl et al., 1993); however, their secondary and tertiary structures differ (Basler et al., 1986; Caughey et al., 1991a; Stahl and Prusiner, 1991; Caughey and Raymond, 1991; Pan et al., 1993; Kocisko et al., 1994, 1995). Many people have resisted the idea that a protein acting alone could cause prion diseases. However, after many years of searching, no one has been able to identify cofactors (such as nucleic acid) or chemical modifications responsible for the conversion of PrPC →PrP Sc (evidence and lack thereof reviewed by Prusiner, 1991 and Caughey and Chesebro, 1997). Therefore, although such factors cannot be ruled out, it seems reasonable to assume that a conformational change in PrPC gives rise to the prion diseases (Caughey et al., 1991a; Baldwin et al., 1995; Harrison et al., 1997; Horiuchi and Caughey, 1999a). In fact, cell-free conversion of PrPC →PrP Sc occurs (in the absence of the GPI anchor and the Nlinked carbohydrates), but the efficiency is relative low (Kocisko et al., 1994). Nonetheless, this cell-free assay conversion of recombinant protein provides strong evidence for the protein-only hypothesis, but demonstration of true infectivity of the converted material is still necessary. Furthermore, DebBurman et al. (1997) have shown that GroEL, a known protein-folding chaperone, increases the efficiency of conversion in the cell-free assay but only in the presence of PrPSc. Difficulties with the purification and production of large amounts of PrP, its tendency to aggregate, and its conformational plasticity have

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prevented the use of x-ray crystallography and multidimensional nuclear magnetic resonance (NMR) spectroscopy for structure determination of full length PrPC and PrPSc until recently, at least for PrPC. Fourier transform infrared (FTIR) and circular dichroism (CD) spectroscopy studies have provided some information (Pan et al., 1993; Safar et al., 1993, 1994). For example, these studies indicate that PrPC is highly helical (42%) with little β-sheet structure (3%). In contrast, PrPSc contains a large amount of β-sheet structure (43%) and less helical structure (30%), is less soluble in aqueous solution, and is resistant to proteolysis. These results, and others, suggest that a conversion of α helices to β sheets may occur on formation of PrPSc from PrPC and that there is a critical core to the molecule (Gasset et al., 1992; Pan et al., 1993; Baldwin et al., 1994; Nguyen et al., 1995; Kocisko et al., 1996). Even given the advances in the NMR studies of PrPC (discussed further later), full characterization of the structural and dynamic attributes of the protein remain elusive. The conformational plasticity of the prion protein has been demonstrated by a variety of techniques for both fragments and the full-length protein. For example, Zhang et al. (1997) have focused on a 142-residue fragment of the Syrian hamster sequence, PrP(90–231), corresponding to the whole of the protein aside from the signal sequence and octarepeat region, which includes all residues involved in the conversion of PrPC →PrP Sc (Fig. 1). This protein adopts two different states rich in α-helical structure. One is found between pH 5–8 and displays NMR chemical shift dispersion and a cooperative thermal unfolding transition consistent with an ordered native state. In contrast, the other helical conformation found at pH 2 has the characteristics of a molten globule or other partially folded state. At elevated temperatures a more stable βsheet-rich state is formed. Similar results were obtained by Swietnicki et al. (1997) for the same fragment with the human sequence. Of note is the fact that the protein undergoes the pH-dependent conformational change in the region of pH 4.4–6. The ∆G° for unfolding at pH 5 is 2.4 kcal/mole (Swietnicki et al., 1997), and the helical content is roughly estimated to drop by approximately 5 to 6 residues based on a change in the ellipticity of 1000 deg cm2 dmol–1 compared with pH 7.2 (Zhang et al., 1997). This pH-induced conformational change has also been observed by Swietnicki et al. (1997) and Hornemann and Glockshuber (1998). Given the pH dependence and that conversion of PrP C → PrP Sc is a posttranslational process that appears to occur in the endocytic pathway, in endosomes or lysosomes (Caughey and Raymond, 1991; Caughey et al., 1991b; Borchelt et al., 1992), lower pH may play a role in facilitating the conformational change that ultimately leads to PrP Sc formation. As with PrPC, conformational heterogeneity is also an issue with PrPSc. For example, various investigators have suggested that PrPSc adopts a

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FIG. 1. Structural features of PrPC. Secondary structure and other structural attributes of the prion protein are labeled along the sequence. The branched symbols in helix B (HB) and the neighboring turn represent the N-linked glycosylation sites. The various fragments used for the NMR studies are indicated by the arrows: NMR1 (Riek et al., 1996); NMR2 (James et al., 1997); NMR3 (Donne et al., 1997); NMR4 (Riek et al., 1997), and NMR5 (Zahn et al., 2000).

variety of conformations and as templates for conversion, these different conformers could explain strain differences (Bolton and Bendheim, 1998; Bessen and Marsh, 1992, 1994; Prusiner, 1991; Bessen et al., 1995; Telling et al., 1996). Direct evidence for strain-dependent structure has been provided by Caughey et al. (1998). They found strain-dependent differences in β-sheet conformation but not in the overall amount of β structure by FTIR spectroscopy. Therefore, the prion protein represents a very challenging system: PrPC contains an ordered domain and a highly mobile region of approximately 100 residues, and PrPSc appears to adopt a variety of different conformations. II. PREDICTIONS OF PrP STRUCTURES AND STUDIES OF PEPTIDE FRAGMENTS TO TEST THE MODELS Until very recently, high-resolution structural information for PrPC was unavailable, and concrete structural information for PrPSc is still

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lacking. As a result, prediction of their conformations was attempted to generate structural models to be tested by experiment. Protein structure prediction remains an unsolved problem, but due to the importance of this system and difficulties getting high-resolution structural information, such modeling efforts were certainly warranted, especially in the way that they were pursued by Cohen, Prusiner and co-workers as described below. Structure prediction was an intimate part of their overall efforts to characterize the properties of the prion protein, such that experimental studies were built around the resulting models so that they could be tested and revised as necessary. A. Prediction of the Structure of PrPC Huang et al. (1994) reported the first predicted structure for PrPC. They used 12 prion protein amino acid sequences (11 mammalian and 1 avian) for multiple sequence alignment, pattern-based turn prediction and a neural network program to predict the secondary structure. The initial secondary structure assignments were ambiguous, so the authors added the constraint that PrP is an all α-helical protein, which was known from the FTIR studies described previously. After assuming that PrP was in the α-structural class, turns and helical segments were identified using a neural net approach (Cohen et al., 1993, Kneller et al., 1990). The results were then mapped onto the sequence alignments, and it was found that there was good conservation of turns, α helices, and the disulfide bonds in the C-terminal region across all sequences. Then, a helix docking routine was utilized to identify good helix-helix pairing sites. These various steps pinpointed four regions suspected to adopt helical structure, and it was predicted that PrPC is a 4-helix bundle protein. Then, a variety of bundle topologies were considered (see Huang et al., 1994 for further discussion). Their preferred bundle, based on a variety of physical criteria, is shown in Fig. 2. The helices are defined as H1, residues 109–122; H2, residues 129–141; H3, residues 178–191; and H4, residues 202–218 (the human numbering system is used). For comparison, Huang and co-workers also investigated three other prediction methods (Chou and Fasman, 1978; Garnier et al., 1978; and the PHD method by Rost and Sander, 1994), where the protein was not designated to be fully α helical. In those cases, the secondary structure predictions for H1–H3 were ambiguous. The Chou-Fasman method predicted H1, H2, and H4 to be helical and H3 to adopt β structure. The Garnier-Robson method was the same except that H2 was also predicted to be a β strand. Finally, PHD predicted that H3 and H4 were helical and H1 and H2 were unknown. Given that H3 and H4 are

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FIG. 2. Predicted structures for PrPC and PrPSc (Huang et al., 1994, 1995). Segments missing from the models are shown as dashed loops.

connected by a disulfide bond and have good helix-helix docking sites, it was assumed that H3 would be constrained to be helical, but hints of the conformationally ambiguous nature of H1 and H2 were revealed. B. Prediction of the Structure of PrPSc The next year Huang et al. (1995) extended their prediction studies to PrPSc. In this case they made use of the conformational ambivalence of H1 and H2 and predicted that they would be the segments of the structure to undergo a conformational transition in the formation of PrPSc. Various packing arrangements were considered, and the preferred model is displayed in Fig. 2 alongside PrPC. This model retains helices H3 and H4, while H1 and H2 are extended and form a β sheet on the helical scaffold. C. Peptide Fragments The predicted structures spurred a number of both experimental and simulation studies to test the predictions and to address the conformational properties of small peptide fragments of PrP. In particular, focus was place on the predicted H1–H4 helices and their conformational properties under a variety of conditions. Although the properties of fragments can often be misleading and have little in common with the properties of the segment in the context of the full-length protein,

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these fragment studies were also performed out of necessity because of the difficulties in obtaining high-resolution information by NMR and xray crystallography. In addition, detailed molecular dynamics simulations in water were performed for the four predicted helical regions for wild-type and mutant sequences corresponding to human-disease-causing variants (Fig. 2, and see DeArmond and Prusiner, 1995). Experimental studies generally yield only limited amounts of information of the actual molecular details of the motion experienced by a peptide or protein, any conformational transitions it undergoes, and the full conformational ensemble. Therefore, there is a need for theoretical studies to better elucidate these events. Complicated macromolecules cannot be studied using analytical methods. Instead, computer simulation methods are used, particularly molecular dynamics, which by providing atomic positions as a function of time, in principle, provides a realistic and complete picture of protein motion. To be most effective, experimental and theoretical studies should be linked. In this way, the experimental results are used to test the validity of the simulations, and when simulations accurately mimic reality they can be used in a predictive manner. Simulations can provide detailed information in these areas that often cannot be obtained experimentally, especially in the case of protein folding where it has proved difficult to characterize proteins during the folding process. Molecular dynamics has become a common technique for simulating the motion of peptides and proteins, and, in fact, it is the most rigorous simulation method available. To use this method, one must have a welldefined starting structure. Another requirement is a potential energy function whose parameters have been derived to reproduce structures and energy trends in various model systems. Using this technique, the atoms move due to their own kinetic energy and the forces exerted on them by all other atoms, including those of the solvent. During the simulation, one can monitor specific interactions both geometrically and energetically to investigate structural transitions and the mechanism by which they occur. Experimentally, the wild-type H1 peptide adopts α-helical structure under various solvent conditions, but it will also quite readily form fibrils composed of β sheets in aqueous solution (Gasset et al., 1992; Zhang et al., 1995; Nguyen et al., 1995). The wild-type H1 sequence maintains a stable helical structure over a 2 ns simulation, as well (Fig. 3, and see Kazmirski et al., 1995). Addition of a valine mutation at residue 117, which is a human-disease-causing mutation (Doh-ura et al., 1989; Hsiao et al., 1991), destabilized the helix and formed extended structure at the C-terminus and may have implications for scrapie formation. Other

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hydrophobic mutations (A117I, A117L, A117NLE, and A117F) did not disrupt the helix (Kazmirski et al., 1995). Other mutations (A117W, A117G, A117P, A117M, A117D, A117N, A117E, A117S, and A117T) were later examined; and A117D, A117N, and A117S formed a similar structure to the A117V peptide with unwinding of the C-terminus (Fig. 3 and

FIG. 3. Molecular dynamics simulations of peptide fragments corresponding to the predicted secondary structure in Fig. 2. H2.5 was discovered while trying to model the missing residues from the PrPC prediction. All simulations began with the peptide in the α-helical conformation, and the average percentage of helix over the 2 ns simulation is given.

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see Kazmirski et al., 1996). Presumably, however, these polar residues would not be tolerated in the hydrophobic core of the protein. Simulations of H2 (now known not to be helical), H3, and H4 (now denoted helices B and C or H2 and H3, Fig. 4), as well as a peptide containing the sequence composing H1–H2 (residues 109–141) were also performed. Simulations of H2 (WT and M129V) starting in a helical state resulted in a loss of helical content (to ~50% helix) but without formation of extended structure nor significant differences between the conformations of the two sequences (Fig. 3). Simulations of H3 (WT and V180I) maintained high helical content (~85%) throughout the simulation (Fig. 3). In contrast, the H4 peptides (WT, V210I, and Q217R) fell to ~50% helical content as a result of the formation of a kink at V209, but the three sequences showed the same effect and the structure did not unwind but merely converted to a helix-kink-helix conformation (Fig. 3). Simulations of the H1–H2 sequence shows that both H1 and H2 can undergo conformational transitions from helix to extended structure and that the Val 117 mutation can trigger loss of helical structure in both H1 and H2. The peptide simulations supported the idea that H1 and H2 may be converted from α helix to β structure, or that H2 may not be helical. Mutations in H3 and H4 yield similar results to WT, suggesting that these mutations may exert their effect on the secondary structure of H1 and H2 in the intact protein and not on the H3 and H4 helices; however, the peptide work is fraught with problems. Small peptides are often conformationally ambivalent and sensitive to the enviroment. H2, H3, and H4 all form helical structure in a helix-promoting solvent (such as hexafluoroisopropanol), yet H3 and H4 readily form β structure when transferred to an aqueous environment (Gasset et al., 1992). Only H2, which is now known not to be helical, retained its helical structure in water. Thus given the ambiguities of working with peptides both experimentally and computationally, simulations of wild-type and mutant sequences in a full three-dimensional model of PrPC were needed, and at the time of the peptide simulations the NMR structures were not available. For that reason, the predicted PrPC structure was revisited. Unfortunately, use of the predicted structure was problematic, as construction of a high-resolution structure based on this model revealed that the orientations of the helices produced a protected core of positively charged residues and no distinct hydrophobic core. In addition, a discontinuous stretch of 20 residues between H2 and H3 was omitted from the model of PrPC of Huang et al. (1994). This segment of the sequence between H2 and H3 is amphipathic with various charged and hydrophobic residues in i → i+4 spacings suggestive of helical struc-

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ture. This sequence was built as an α helix in the same manner as H1–H4 and found to maintain a stable helical conformation through 2 ns of MD (H2.5, Fig. 3). In summary, wild-type and all human-disease-causing mutant fragments of H1–H4 were simulated for at least 2 ns in water beginning in a helical conformation. The results suggested that the effects of the mutations must also be evaluated in the context of the rest of the protein, (i.e., in a model of PrPC). Simulations using the four-helix bundle structure of Huang et al. (1994) were attempted; however, there were several errors in the model, and it was deemed to be chemically and physically flawed, aside from the last two helices. For that reason, further secondary structure prediction was performed using 11 different structure prediction algorithms, including the 4 used previously, and 23 sequences for the multiple sequence alignment (Alonso et al., 1996). These efforts and the fragment simulations described previously led to the prediction that there are two small extended strands in PrPC and another helix (H2.5 between H2 and H3, Fig. 4 and Alonso et al., 1996). For unknown reasons, not all of the results of the original prediction were reproduced even by using the same algorithms (methods 4, 6, and 8–10 in Fig. 4). Fortunately the NMR structure by Wüthrich and co-workers came out not long after this and verified the predicted strands, and the

FIG. 4. Comparison of secondary structure prediction results from a variety of methods for residues 90–231 of PrP. The original prediction is that of Huang et al. (1994). The NMR assignments are those of Riek et al. (1996) and James et al. (1997). The new predictions were made considering the results of 11 different secondary structure prediction algorithms. Although H3 was predicted to adopt both extended and helical conformations, we reasoned that H3 was helical and constrained by the disulfide bond to H4. H3 is indeed helical in the new NMR structures. H2.5 was first identified from the sequence and tested with MD. These more in-depth predictions support the idea that it is helical, and it has also been found by NMR. The status of H1 is still not clear. There are some indications by NMR that it adopts some weak, marginally populated helical structure. These methods used are numbered 1–11 and correspond to the following: (1) Consensus prediction from the IBCP Server (http://www.ibcp.fr) consisting of methods by Geourjon and Deleage (1994), Gibrat et al. (1987), Levin et al. (1986), Deleage and Roux (1987), and Rost and Sander (1994); (2) SOPMA, Geourjon and Deleage (1994) from IBCP Server (http://www.ibcp.fr); (3) Levin et al. (1986) from by IBCP Server (http://www.ibcp.fr); (4) PHD, Rost and Sander (1994), EMBL Server (http://www.embl-heidelberg.de/predictprotein/phd_pred.html) (5) Gibrat et al. (1987) from IBCP Server (http://www.ibcp.fr); (6) enhanced neural network, Kneller et al. (1990), no secondary structure constraints; (7) White et al. (1994) from PSA Server (http://bmerc-www.bu.edu/psa/); (8) Chou and Fasman (1978); (9) enhanced neural network, Kneller et al. (1990), helical protein, UCSF server (http://www.cmpharm.ucsf. edu/~nomi/nnpredict.html) (10) Garnier et al. (1978) no secondary structure constraints; and (11) Garnier et al. (1978) 20–50% Helical constraint.

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new helix and revealed the overall topology of the molecule, which could not be predicted with any confidence. III. PrPC STRUCTURAL MODELS FROM NMR The first NMR structure of a relatively large PrP fragment became available in 1997. The structure was determined by Wüthrich and coworkers (Riek et al., 1996) for the murine PrP fragment containing residues 121–231 (Fig. 1), PrP(121–231). This study, and subsequent NMR work, represented a tremendous advance in the area of prion research, in general, and in the area of prion modeling and computational studies, in particular. At the time, the structure was rightly criticized as being potentially irrelevant by virtue of being a fragment, as previous studies demonstrated that smaller fragments of PrP adopt a variety of conformations depending on solution conditions. However, the new structures reported in 1997 confirmed the overall structural features of the PrP(121–231) fragment (Fig. 1) and supported the contention that these residues constitute an autonomous folding unit. One of the newer structures is of Syrian hamster PrP(90–231) (James et al., 1997). The other two structures are full-length versions of these two fragments, comprising all residues including the N-terminal octarepeats (Riek et al., 1997; Donne et al., 1997; Fig. 1). Also, more recently, James and co-workers followed up their work on Syrian hamster (90–231) and presented more new data and the refined structure (Liu et al., 1999). The two primary PrP fragments on which the structures for the fulllength proteins are based are displayed in Fig. 5. The overall folds of the two average NMR structures are similar, as would be expected for sequences exhibiting > 90% homology. They contain three wellresolved helices and a short two-stranded β sheet. Differences are evident: helix B (H3 above) is 6 residues longer and helix C (H4 above) is 10 residues longer in the James’ structure (Fig. 5). This widespread increase in helical content is consistent with the pH-dependent conformational behavior described previously; the Riek et al. (1996) mouse structure was determined at pH 4.5 in water and that of the hamster protein by James et al. (1997) was at pH 5.0 in acetate buffer. In addition, however, the differences could be caused by the differences in the sequence or the lengths of the fragments. The short and slightly variable β sheet is similar in the two structures and corresponds to the regions predicted in Fig. 4. In addition helix A corresponds to the new predicted helix in Fig. 3 and 4 (H2.5). In contrast, the loop between S2 and helix B (Fig. 5) is disordered in the smaller fragment but gives rise

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to many medium- and long-range NOEs (nuclear Overhauser effect cross-peaks indicating that protons are near in space) in the longer protein. Thus, overall the structures are very similar and would appear to be part of a continuum, with the James et al. conformation being more structured and presumably closer to the physiological form of PrPC. An obvious difference between the two fragments is the N-terminal extension on the James et al. structure (Fig. 5). This region is particularly interesting because it is implicated in the conversion of PrPC → PrPSc. It is conformationally heterogeneous and overall disordered by NMR, but note that both of these fragments were studied at a pH falling

FIG. 5. Ribbon representations of NMR1 (mouse), NMR2 (Syrian hamster), and NMR5 (human) (Fig. 1). The dark portions of the helices correspond to helical extensions relative to NMR1.

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within the pH-dependent conformational transition described previously and also the protein undergoes rapid interconversion between a weak dimer and monomer (James et al., 1997). As such, even the longer fragment may not be entirely representative of the conformation under physiological conditions. Part of this region (residues 109–121, H1) was predicted to be helical (Gasset et al., 1992), and it does indeed form a helix under different solvent conditions (Gasset et al., 1992; Nguyen et al., 1995; Zhang et al., 1995). Furthermore, the last four residues of the predicted H1 helix display only moderate exchange of their amide hydrogens and the α-proton and α-carbon chemical shifts for residues 90–127 are consistent with weak α-helical content. There are nine nonsequential NOEs in this region, implying that some structure exists at least transiently. However, this is in marked contrast to the other three well-ordered helices. In any case, the work from James group provides some evidence for weak helical structure in this region of the protein and structural information for the protein at higher pH is needed. Also, it is worth noting that even outside the H1 region, these NMR structures should be viewed as low-resolution structures. For example, the James et al. structures (NMR2) have an average of 2.5 (or ~3 in the more structured region of the protein) unambiguous, long-range NOEs per residue. Both NMR groups have followed up on their fragment studies with experiments on the full-length PrP protein, PrP(23–231) or PrP(29–231) (Riek et al., 1997; Donne et al., 1997). In essence, these investigations have confirmed the previous studies: The core of the molecule composed of the three helices and short β sheet is retained, and the rest of the molecule is disordered and/or highly mobile. New structures, per se, were not determined, and the flexible N-terminus is represented schematically on the framework provided by the fragment studies. Although these studies have provided little new structural data, they are important controls because one must always question the relevance of data obtained from fragments. The most striking new result of these studies is the marked difference in flexibility and dynamic behavior between the N-terminus and the folded core of the molecule, as probed by the heteronuclear 15N-1H NOE and described by the related 15N-1H correlation time. These studies conclusively demonstrate the plasticity of the N-terminal region of the prion protein. More recently a NMR structure for a fragment of the human form has been determined by Wüthrich and co-workers (Zahn et al., 2000; Fig. 5). Despite the NMR structures now available, modeling and simulations studies are still necessary for the following reasons: (1) both PrPC and PrPSc are conformationally heterogeneous, (2) the ‘PrPC’ structures were

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determined under conditions where the protein undergoes the pHinduced conformational change to scrapie, and (3) NMR experiments, and almost all other experimental techniques, report on the average properties of the protein ensemble. In addition, there is little to no structural information about the conversion process and subsequent aggregation. Thus, although not ideal, theoretical and computational studies represent a logical avenue for investigating the structural properties and conformational transitions of this system.

IV. DETAILED MODELING STUDIES BASED ON THE NMR MODELS With the various PrPC NMR models we can now begin to address possible effects of mutation, causes of species barriers, and strain differences, and the mechanism of conversion of PrPC → PrPSc. We are still plagued, however, by the lack of high-resolution structural information for PrPSc. The theoretical/modeling studies fall into two groups: (1) those involving analysis of the static NMR structures to rationalize the known experimental data and to formulate hypotheses and (2) those that use the NMR models as starting structures for detailed computer simulations to investigate the dynamic characteristics of the protein and the conversion of PrPC → PrPSc. A. Static Modeling 1. Overall Features of the Models The three independent NMR structures now available for the mouse, Syrian hamster, and human sequences are all quite similar (Fig. 5). Some of the overall features of the models will now be discussed, using the new human structure for the purpose of illustration. Unfortunately, this structure begins at residue 125 and ends at 228. Thus, coordinates for residues 90–124 and 228–232 from the hamster structure were used, and the appropriate sequence changes were made to yield the structure in Fig. 6A. The 3F4 and 13A5 antibody binding sites are shown in Fig. 6B, in the same orientation. In addition, other antibody binding sites have been generated, and they cluster into three regions, termed A-C (Peretz et al., 1997), and these are shown in Fig. 6C (again in the same orientation). PrP-directed antibody studies have also produced a PrPSc specific antibody (Korth et al., 1997). This antibody recognizes a discontinuous epitope, which could form a continuous surface after a conformational transition or on aggregation (Fig. 6D). The region involved

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in the binding of PrPSc that precedes the conformational change (Horiuchi and Caughey, 1999b) is shown in Fig. 6E. Antibodies to the highlighted area on PrPC in Fig. 6E prevent interactions with PrPSc by blocking a binding site nearby. Fig. 6F shows the model with Asn-linked oligosaccharides to illustrate the relative sizes of the protein and its attached carbohydrates (see DeArmond et al., 1999 for further discussion of the modeling of the carbohydrate groups). (Note that this panel is not to scale with respect to the others, and it is has a different orientation). Of interest, there are many positively charged residues lacking salt-bridging partners on the bottom surface of the molecule, as depicted in Fig. 6F, and adjacent to the C-terminus, which is the position of the GPI anchor to the membrane. Thus, the residues appear to be positioned to interact with the phosphate groups on the membrane. 2. Species Barriers Billeter et al. (1997) explored the structural basis of the species barrier for transmission of prion diseases. They showed that the variability of 31 mammalian PrP sequences maps onto three distinct regions of their NMR structure of PrP(121–231). One of these (class A) is the putative protein X binding site (the putative protein X binding site overlaps and extends the large patch of residues at the bottom right in Fig. 6D). The sequence variability in this region involves changes in the electrostatic charge of the side chains, which may alter protein-protein recognition and binding. Another is near the proposed PrPSc binding site (class C) and again involves polar residues that may be involved in recognition and binding (Fig. 6E). Finally, class B overlaps with class C FIG. 6. Structural properties of the human PrPC NMR structure. (A) The main chain fold of NMR5 but modeled with extensions from the Syrian hamster structure to yield HrPrP(90–232). The secondary structure is colored gray, using the assignments of Zahn et al. (2000): S1, residues 128–131; HA, 144–154; S2, 161–164; HB, 173–194; and HC, 200–228. (B) A space filling version of panel A with the 13A5 (residues 138–141) and 3F4 (residues 106–115) epitopes highlighted. The protein is in exactly the same orientation as in panel A. (C) A space filling representation of model with various antibody binding epitopes displayed: epitope A = residues 96–105, B = 152–163, C = 225–231 (Peretz et al., 1997). (D) The discontinuous PrPSc-specific antibody binding site (Korth et al., 1997) is displayed (residues 142–148, 162–170, 214–226). (E) Antibodies to the highlighted region (residues 219–232) on PrPC prevent binding of PrPSc, although it appears to be due to steric blocking of a nearby binding site, not to binding specifically in the region indicated (Horiuchi and Caughey, 1999b). (F) N-linked carbohydrates at N181 and N197 are displayed to indicate the relative bulk of the sugars and the regions of the molecule “protected” by them (DeArmond et al., 1999). Note that the scale and orientation are different than in the other panels. Also, the Lys, Arg, and His residues are displayed. In particular, the positively charged residues at the bottom of the molecule are predicted to interact with the phosphate head groups of the membrane.

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but is composed of buried hydrophobic residues, whose involvement in determining the species barriers are presumably manifested after a conformational change that would bring about their exposure. Warwicker (1997a) has extended Prusiner’s proposal that a PrPC/PrPSc heterodimer could be a key intermediate in the infectious process (Prusiner, 1991) and suggests that the dimer interface is involved in normal physiological events (for example, as PrPC/PrPC) and that scrapie merely exploits a normal recognition surface for pathological consequences. This hypothesis led him to search for conserved regions of the sequence in an attempt to identify the “normal” binding surface. The species variations are then considered in light of this putative homodimer recognition surface. He focused on eight of the better studied residues involved in species barriers, and they fall either within or near this binding surface. 3. Strain Differences A related issue is strain differences with respect to PrPSc. Strains in the absence of genetic material are of course peculiar. Strains have been explained in terms of multiple conformations of PrPSc. In this model PrPSc serves as a template for conversion such that conformational diversity leads to different forms of endogenous newly converted PrPSc (Telling et al., 1996). Studies by Caughey et al. (1998) provide direct evidence for strain-dependent difference in the β-sheet conformations by FTIR. Interestingly, the differences are in the type of β structure as opposed to the overall amount or proportion of β structure in the protein. Given that PrPSc structures or even sound structural models are not yet available, details regarding the nature of the conformational heterogeneity are lacking. Another factor besides PrPSc conformations in strain differences could be differences in PrPC/PrPSc interaction surfaces, as with the models for species barriers discussed previously. For example, Warwicker (1997b) has interpreted his heterodimer model in light of residues important in determining strain differences. In particular, in this case, he focused on conserved and nonpolar residues and proposed that there are two distinct patches on the molecule that can interact with membrane or with a neighboring protein. This hypothesis leads to two different membrane-attached PrP orientations, or faces, that would be presented to an incoming PrPSc molecule. In a more recent study, Warwicker (1999) discusses how charge interactions between PrP and the membrane may affect scrapie formation. Along these lines, Morillas et al. (1999) have reported that there is significant ordering of the N-terminus of the human protein on association with membranes, and the C-ter-

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minal, folded domain becomes destabilized on binding to the membrane surface. Despite the heterogeneity that could result from the mechanisms described previously, the strain behavior is difficult to explain entirely by a phenomenon involving protein-protein interactions or multiple conformations of PrPSc. However, there is certainly evidence supporting this idea (Telling et al., 1996; Kocisko et al., 1994, 1995; Bessen et al., 1995). PrP is a glycoprotein and the carbohydrate groups may add another level of complexity to increase the diversity of PrPSc, as it is estimated that 401 different PrP glycoforms are possible (Endo et al., 1989). PrPSc molecules from different prion strains differ in size and state of glycosylation (Stahl et al., 1993), and there can be differences in glycosylation of PrPC versus PrPSc (Rudd et al., 1999). DeArmond et al. (1997) have shown that different regions of the brain synthesize their own unique sets of PrPC glycoforms and that homologous regions in hamsters and mice express animal species-specific PrPC glycoforms. Furthermore, more recent studies by DeArmond and et al. (1999) demonstrate that the selective targeting of neuronal populations is determined by cell-specific differences in the affinity of an infecting PrPSc for PrPC and that this affinity can be modulated by nerve-cell-specific differences in PrPC glycosylation. Thus, the carbohydrate groups might influence PrPSc/PrPC interactions, and therefore replication of PrPSc. They may do this through direct interactions involving protein recognition events, which are reasonable given the way in which the carbohydrates change the surface displayed by the protein (Fig. 6F), or through modulation of the structure, dynamics, and stability of PrP. Thus, it appears that there are sufficient animal species-determined variations in the amino acid sequence of PrPC, the level of expression of PrPC, and the carbohydrate structure of PrPC in combination with variations in the conformation and amino acid sequence of PrPSc to account for the known variations in the prion disease phenotype and strain differences in the absence of genetic material. Unfortunately, more detailed structural information about PrPSc and heterodimers is necessary for more rigorous modeling of the possible factors involved in strain differences. 4. Human Disease-Causing Mutations The NMR spectroscopists and other modelers have explored the possible structural basis for the familial prion diseases. The premise is that inherited prion diseases are due to destabilization of PrPC on mutation, which facilitates PrPSc production. The known human disease causing mutations (Fig. 1) have been mapped onto the extended human NMR

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structure (with the N-terminal residues 90–124 residues from James et al., 1997 included) in Fig. 7. The mutations span the full sequence of the protein and occur in various types of structure. Also, they occur both at the surface and in the hydrophobic core. In fact, it is difficult to rationalize the effects of these mutations based solely on the structure of PrPC. Nonetheless, Riek et al. (1998) have discussed the possible structural ramifications of the nine disease-causing mutations in the structured position of the molecule (Fig. 1). For example, the T183A mutation would eliminate two hydrogen bonds in the hydrophobic core that link helix B and the β sheet (Fig. 7B). D178 forms a salt bridge with Arg 164 and a hydrogen bond with the hydroxyl group of Y169 (Fig. 7C). Riek et al. (1998) predicted that these interactions would be disrupted on mutation of D178 → N and reduce the thermodynamic stability of the mutant relative to wild-type. This site is also interesting because the effects of this mutation are linked to the polymorphism site, Met129/Val129, and Riek et al. proposed that the hydrogen bonding network around residue 178 is affected by which residue is at position 129. The Q217R mutant was predicted to be destabilizing, as Gln is surrounded by hydrophobic groups, and it forms hydrogen bonds with a main chain carbonyl. The F198S mutation introduces a polar residue into the hydrophobic core, as well as introducing a cavity. These effects are predicted to destabilize the protein. The other four mutations— V180I, E200K, R208H, V210I—were predicted to be well tolerated with respect to structure and stability. Swietnicki et al. (1998) evaluated the secondary structure and thermodynamic stability of the P102L and E200K disease-causing mutants and found that the mutations had little effect on the structure and stability of the protein, which is consistent with the prediction regarding E200K. Recent experimental work by Liemann and Glockshuber (1999) addressed the thermodynamic stability of the disease-causing mutations discussed by Riek et al. (1998). They found, that, overall, the mutations do not lower the stability of PrPC appreciably at neutral pH, and they concluded that destabilization on mutation is unlikely to be a determinant for the PrPC → PrPSc conversion. Nevertheless, there is decent qualitative agreement between the predictions of Riek et al. (1998) and the experimental values. Aside from T183A (destabilized by 4.6 kcal/mole relative to wild-type), however, the values are low (7 of the 9 mutants—including the polymorphism at residue 129—are destabilized by ~2 or less kcal/mole) (Liemann and Glockshuber, 1999). In addition, contrary to the predictions, the D178N/M129 and D178N/V129 proteins have similar properties and are destablilized to the same extent (1.7 and 1.9 kcal/ mole, respectively).

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FIG. 7. Positions of the human disease causing mutations in the human NMR structure (Zahn et al., 2000). (A) The mutations listed in Figure 1 are highlighted. (B) Close up of the T183 interactions, which form solvent excluded hydrogen bonds connecting HB and S2. (C) Close up of D178 salt bridge and hydrogen bonding interactions on the surface that also connect HB and S2.

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In contrast, Cappai et al. (1999) reported a decrease in helical structure and altered thermal stability in the case of the P101L mutation, and more importantly they see that stability is sensitive to pH. In fact, the structural differences between the P101L and wild-type proteins increased both above and below pH 7. It would be helpful to see Liemann and Glockshuber’s studies performed as a function of pH to rule out differences in stability at the lower pH values presumed to be important in the conversion process. Also, it should be noted that the experiments were performed on different sizes of PrP, different sequences (human and mouse); and all were done in the absence of carbohydrate groups, copper ions, and the GPI anchor, all of which can affect the structure and stability of the protein. Nonetheless, it may well be that the mutations do not cause a change in thermodynamic stability and instead lower the energy of the transition state for the conformational change and thereby accelerate conversion. More details regarding the kinetics and molecular motions involved in the conversion to PrPSc are necessary to evaluate these alternatives. In this regard, atomic-level molecular dynamics simulations of wild-type and mutant forms of the protein may begin to provide a structural framework for familial prion diseases. B. Detailed Atomic-Level Molecular Dynamics Simulations of PrP 1. PrPC Simulations Two molecular dynamics simulation studies of PrPC (neutral pH) have recently been reported. In the first, Zuegg and Gready (1999) began with the Syrian hamster structure (NMR2) of James et al. (1997; Liu et al., 1999). They also constructed and simulated a human homology model. Their simulations are short (< 1 ns), so it is unlikely that they have equilibrated. They found that the structures were very unstable and that the helices began to unravel. By ~720 ps the main chain root-mean-square (RMS) deviation was 5.5 Å. They attributed the problems to using a short (8 Å) nonbonded cutoff (electrostatic and van der Waals interactions are only evaluated within a certain range to decrease the computational cost). However, the same cut-off and starting structure were used below, and the RMS deviation was only ~1.7 Å at 720 ps (0.72 ns) and in fact remains ~2 Å over the 10 ns of the simulation. It would appear that there are other problems with these simulations. Zuegg and Gready were able to obtain better results by using a method to account for longer range electrostatic interactions, but the other problems still remain and in particular quite nonstandard simulation protocols were used.

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Much more standard protocols were used by Parchment and Essex (2000) in their simulations of the mouse (NMR1) and hamster (NMR2) proteins. Their simulations, particularly that of the hamster, had not yet equilibrated by the end of the 1 ns, which makes the conclusions a bit tentative. Nevertheless, their simulations were much more stable and the proteins were better behaved. By comparing the simulations of MoPrP(121–231) and Sha(113–231), they found that the extra residues at the N-terminus seem to affect the dynamics and stability of the secondary structure. In particular the β sheet is longer in the hamster simulation, but it is less stable. Also, the helices are stable in both simulations, but the rest of the structure was quite dynamic in the case of the hamster protein. The mouse protein was not as dynamic. Such differences in the behavior of the two proteins is interesting given that there are only six amino acid changes. 2. Simulations of the Conversion of PrPC → PrPSc In light of what is known about the prion protein and its fragments, the working hypothesis is that the conversion of PrPC → PrPSc is due solely to a conformational change in the protein, such that the two forms are conformers. Safar et al. (1994) suggested that the scrapie form is an aggregate from a molten globule folding intermediate, and as described previously, the lower pH of the endosomal pathway appears to trigger a conformational change to PrPSc or something intermediate between PrPC and PrPSc. The experimental studies thus far provide evidence for a conformational change linking PrPC and PrPSc, but localization of the change along the sequence has proved difficult. Peretz et al. (1997) have produced antibodies to the N-terminal region (residues 90–120) that recognize PrPC but do not bind PrPSc (region A in Fig. 6C). In contrast, an epitope at the C-terminus is accessible in both forms. The authors argue that the major conformational change occurs at the N-terminus, which is consistent with the plasticity of this region, but it is possibile that the N-terminal epitope does not change conformation and instead no longer binds because the surface is unavailable in the aggregate. However, it is unlikely that PrPC would retain a disordered Nterminus in PrPSc, as even amorphous aggregates typically display specific packing arrangements (Speed et al., 1996; Fink, 1998). Also, the PrPSc specific antibody (Korth et al., 1997) may provide hints about the structure of PrPSc and/or its aggregation properties. In support of the idea that the N-terminal region is important in the conversion, it was found that peptide fragments from the 119–140 region can inhibit the conversion reaction (Chabry et al., 1998, 1999). These various low-

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resolution clues about conversion can be used to evaluate simulations in which a conformational change is triggered by lowering the pH. Molecular dynamics simulations beginning from the Syrian hamster NMR structure of James et al. (1997) (residues 109–219) have been performed at low (Asp, Glu, and His residues protonated) and neutral/high pH (above the pKa of His, His neutral) in water for 10 ns each (Alonso et al., in press). The disulfide bond was kept intact, as evidence points to it remaining oxidized in authentic PrPSc and necessary for infectivity (Turk et al., 1988; Somerville et al., 1980; Hermann and Caughey, 1998). It is not clear if the β-sheet-rich states obtained with the reduced protein are relevant (Jackson et al., 1999a, 1999b). The hypothesis for these simulations is that the protein will be stable at the high pH, but will undergo a conformational change at the low pH. It should be noted that our expectations regarding stability are not overly strict for this system because of the pH of the NMR experiments and the low number of long-range NOEs, which yield fairly low resolution structures (2.5 unambiguous long-range NOEs per residue compared with a more typical value of four to five such long-range contacts in well-behaved proteins). Although a lack of NOEs can be due to a variety of factors, heightened mobility appears to be the primary cause based on NMR relaxation experiments (Donne et al., 1997; Riek et al., 1997). All of this makes precise determination of a unique structure difficult, and even in cases where one has many more long-range NOEs, different structure-generating protocols and programs can yield different “solution structures” from the same NMR data, suggesting that the actual family of NMR structures may be larger than indicated on the basis of calculated structures using a single method (Liu et al., 1992). Furthermore, for a given density of local NOE restraints, NMR structuregenerating methods have their own biases in conformational sampling and ultimately the structures they produce. Because of these issues, we concentrate on fairly large comparative conformational changes. In any case, the Cα RMS deviations for the high pH simulation do indeed remain relatively low, particularly for the core of the molecule (i.e., disregarding the N-terminus), whereas the structure deviates more dramatically at low pH (Fig. 8A). The heightened mobility of the N-terminal region is evident in the Cα RMS fluctuations about the MD mean structure over the last 2.5 ns of each simulation (Fig. 8B). At high pH, the fluctuations about the secondary structure are low, and the only high values correspond to the N-terminus and the loop between HB and HC. Various snapshots from the two simulations are overlaid in Fig. 9 to illustrate the nature and extent of the motion. The low, and quite normal, mobility of the structured part of the molecule is in agreement

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FIG. 8. Simulations of PrPC at low and neutral/high pH. (top) The root-mean-square (RMS) deviation from the starting NMR structure (NMR2) as a function of simulation time. (Bottom) The RMS fluctuation of the α carbons about their mean structure during the molecular dynamics simulations following equilibration (over the last 2 ns).

with hydrogen exchange studies of the human protein by Hosszu et al. (1999). In contrast, high mobility is observed throughout the sequence at low pH, and both the motion and deviations from the starting structure for the first 70 residues (including the N-terminus, S1, HA, and S2) are striking (Fig. 8B). Examination of structures as a function of time indicates that at high pH the protein mostly just fluctuates about a mean structure that is very similar to the starting structure (Fig. 9). In contrast, at low pH there are motions spanning many Ångstroms in the N-terminal half of the mole-

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FIG. 9. The main chain fold at various time points during the neutral/high pH and low simulations PrP. The protein is dynamic, particularly in the turns and loop regions, but the overall structure is well maintained at neutral/high pH. In contrast, at low pH the conformational transitions and fluidity span almost the entire structure.

cule. The C-terminal helices are stable. The S1 and S2 strands constitute a very short β sheet in the starting structure (3 residues each). However, in the simulation at low pH both strands elongate to lengthen the sheet. Also, the neighboring N-terminus was brought into the sheet via hydrophobic contacts between side chains. These changes resulted in growth of the sheet in both length and width (Fig. 10). Residues 121–124 are interesting because they are consistently predicted to form β structure (Fig. 4). The sheet propagated in this region and involved contacts between residues 128 and 165. However, the extended structure is distorted and dynamic and lacks some of the precise packing and hydrogen bonds one would like to see in β structure. In addition, HA and its preceding loop moved by approximately 10 Å over the course of the simulation to bring HA into closer proximity to the helix scaffold. Also, HA experienced some loss of helix at its C-terminus to accommodate the movement. The loop appears as if it is moving up to join S2 (Fig. 10). Further consolidation of the structure may occur with time, or, alternatively, it is possible that the precise β structure is obtained only on dimer formation with another PrPSc molecule. Why does PrPC have the potential to undergo this conversion? Several features mark the region around N-terminal region, S1, and S2 as potential triggers for conversion to β structure. The N-terminal region is conformationally heterogeneous and is able to easily adopt both helical and

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FIG. 10. Time progression of the simulation of PrPC at low pH, conditions supporting conformational change to PrPSc. Extension of the β sheet occurs and other strands are brought into the sheet.

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more extended conformations. Many residues in this region are already extended in PrPC but are not actually participating in secondary structural contacts (for example, residues 120–129, 131–144, 155–161, 165, 166). Furthermore, residues 121–124 and 136–141 have the propensity to form β structure (Fig. 4). The short existing β sheet is in close proximity to these residues and appears to serve as a structural nucleus for the formation of new and propagation of existing β-like structure. Although not definitive, the results of the molecular dynamics simulations at low pH suggest that this approach may provide insight into the early steps in the conversion of the helix-rich state of PrPC to PrPSc, a conformer with increased β-structure content and diminished helical structure. The simulation of the isolated monomer reflects the tendency of the N-terminal portion of the sequence to convert to β-likestructure, but the structure is still loose and would appear to require intermolecular interactions with another molecule to obtain further consolidation of the β structure. As such, the simulations may be capturing the early steps in the pH-catalyzed transformation to PrP* or an intermediate in route to PrPSc. Acid-induced unfolding intermediates have been observed by Swietnicki et al. (1997) for a human fragment of residues 90–231. It was assumed that the bulk of the change in structure was in the N-terminal region of the protein. However, Hornemann and Glockshuber (1998) have also observed a scrapie-like unfolding intermediate of PrP(121–231). So, although the most dramatic changes in the low pH simulation occurred in the N-terminal region of the protein, the other changes in the C-terminal portion are also supported by experiment. With a detailed model of PrPSc, or something approaching PrPSc, we are now in a position to investigate possible binding interactions between PrPC and PrPSc, to build testable models of PrPSc aggregates, and to attempt rational drug design. ACKNOWLEDGMENTS We are grateful for financial support provide by the National Institutes of Health (GM50789). UCSF Midas Plus (Ferrin et al., 1988) was used to make Figures 3, 6, 7, and 9. Molscript (Kraulis, 1991) was used to make Figures 2, 5, and 10.

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INTERACTIONS AND CONVERSIONS OF PRION PROTEIN ISOFORMS BY BYRON CAUGHEY, GREGORY J. RAYMOND, MICHAEL A. CALLAHAN, CAI’NE WONG, GERALD S. BARON, AND LIANG-WEN XIONG Laboratory of Persistent Viral Diseases, NIAID, NIH, Rocky Mountain Laboratories, Hamilton, Montana 59840

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. PrP and Transmissible Spongiform Encephalopathies . . . . . . . . . . . . . . . B. Nomenclature of PrP Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. TSE-associated Changes in PrP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Folding and Unfolding of PrP-res . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Formation of PrP-res by Deleted Forms of PrP . . . . . . . . . . . . . . . . . . . . . C. Structural Diversity in Abnormal TSE-associated PrP Molecules . . . . . . . III. Mechanistic Models of PrP-res Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Binding Interactions between PrP-sen and PrP-res . . . . . . . . . . . . . . . . . . . . V. PrPSc-induced Conversion of PrP-sen to PrP-res: Biological Connections . . VI. Mechanistic Studies of the PrP-res-induced Conversion Reaction . . . . . . . . A. Importance of the Disulfide Bond . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Sites of Interaction between PrP-sen and PrP-res . . . . . . . . . . . . . . . . . . . VII. PrP-sen/PrP-res Interactions and Species Barriers. . . . . . . . . . . . . . . . . . . . . A. Conversion Efficiency versus Susceptibility . . . . . . . . . . . . . . . . . . . . . . . . B. Chronic Wasting Disease. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Binding versus Conversion of Heterologous PrP Molecules . . . . . . . . . . VIII. TSE Strains and PrP-sen/PrP-res Interactions . . . . . . . . . . . . . . . . . . . . . . . . IX. PrP Perturbations and TSE Infectivity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . X. PrP-sen/PrP-res Interactions and the Search for Anti-TSE Drugs. . . . . . . . . XI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

139 139 140 141 141 145 145 146 148 150 153 153 154 156 156 157 157 159 162 162 164 164

I. INTRODUCTION A. PrP and Transmissible Spongiform Encephalopathies The odd properties of the infectious agents of the transmissible spongiform encephalopathies (TSE or prion diseases) evoked proposals that they might represent a distinct class of infectious agents (often called prions), which are proteinaceous and devoid of nucleic acid (Griffith, 1967; Prusiner, 1982; Bolton and Bendheim, 1988; Prusiner, 1998). A key ingredient to these proposals is that the infectious agent is an abnormal form of a host-encoded protein that can interact with its normal counterpart and cause it, too, to become abnormal. Considerable evidence suggests that prion protein (PrP) is the host protein that, 139

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when corrupted, is the culprit in TSE diseases (reviewed in Prusiner, 1998; Horiuchi and Caughey, 1999). The gene for PrP is clearly essential for transmission and pathogenesis (reviewed in Prusiner, 1998; Horiuchi and Caughey, 1999). The “protein-only” or prion hypothesis asserts that an abnormal form of PrP, probably the aggregated, amyloidogenic protease-resistant form (PrP-res or PrPSc), is the infectious agent that propagates itself by inducing a change in the normal, protease-sensitive PrP (PrP-sen or PrPC) (Prusiner, 1998; Horiuchi and Caughey, 1999). Indeed, PrP-res usually correlates with TSE infectivity (Prusiner, 1998; Horiuchi and Caughey, 1999) and can induce PrP-sen to convert to the protease-resistant, aggregated state in cell-free reactions (Kocisko et al., 1994). These conversion reactions are highly specific in ways that reflect, and may explain, important aspects of TSE biology (see later) (Horiuchi and Caughey, 1999). However, although considerable progress has been made in understanding the structures (Wüthrich and Riek, this volume) and folding options (Glockshuber et al., this volume) of various PrP molecules, uncertainties remain about the mechanism of PrP-res formation and its relationship to TSE infectivity (Chesebro, 1998; Caughey, 2000). This chapter focuses on the structure(s) of PrP-res, its interactions with PrP-sen, and how such interactions appear to relate to the propagation of TSE agents, the transmissibility of TSE agents between species, the existence of TSE strains, and the development of TSE therapies. B. Nomenclature of PrP Isoforms PrP is normally a protease-sensitive sialoglycoprotein that is anchored to the plasma membrane via glycosyl-phosphatidylinositol (GPI) (reviewed in Caughey and Chesebro, 1997; Prusiner, 1998; Weissmann, 1999 and Priola and Baldwin chapters, this volume). We will use the term PrPC to refer to PrP in its normal structure and conformation and the term PrP-sen to refer generically to protease-sensitive forms of PrP, whether normal (i.e., PrPC) or not (e.g., various recombinant forms). The terms PrPSc, PrPCJD, PrPBSE, etc., refer to abnormal forms of PrP associated with the particular TSE disease, and depending on the individual usage, may variably imply one or more of the following properties: infectiousness, pathogenicity, protease-resistance, or a specific conformational state. More generically and operationally, we refer collectively to protease-resistant forms of PrP as PrP-res. However, the structures and properties of various abnormal PrP molecules are diverse, making the precise definitions and use of these terms problematic (see Preface and later). Nonetheless, with that caveat, we will use

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the generic term PrP-res, unless referring specifically to the abnormal PrP forms associated with a particular TSE disease. II. TSE-ASSOCIATED CHANGES IN PrP In most TSE diseases, the same host gene encodes both PrPC and PrP-res. In scrapie-infected neuroblastoma cells, at least, PrPSc is made from mature PrPC (Borchelt et al., 1990) after it reaches the plasma membrane (Caughey and Raymond, 1991) (see Harris, this volume). The conversion site is likely to be either the cell surface and/or along an endocytic pathway to lysosomes (Caughey and Raymond, 1991; Caughey et al., 1991; Borchelt et al., 1992). PrPC and PrPSc appear not to differ regularly in covalent structure (reviewed by Baldwin, this volume); however they can be discriminated in several other ways (reviewed in Caughey and Chesebro, 1997; Prusiner, 1998; Weissmann, 1999). The rates of biosynthesis and turnover of PrPSc are much slower than those of PrPC (Caughey et al., 1989; Borchelt et al., 1990). PrPC is fully digested by proteinase K (PK), whereas PK usually removes only roughly 67 of the ~210 total residues from the N-terminus of most PrPres (PrPSc and PrPCJD) molecules (Oesch et al., 1985; Hope et al., 1986; Hope et al., 1988). PrPC is soluble in mild detergents, whereas PrP-res is much less soluble and tends to assemble into amorphous aggregates or amyloid fibril-like structures (scrapie-associated fibrils or prion rods) (Merz et al., 1981; Diringer et al., 1983; Prusiner et al., 1983). A. Folding and Unfolding of PrP-res Spectroscopic studies have determined the high-resolution structure of recombinant PrP-sen and shown that it has high α-helical content but little β sheet (Pan et al., 1993; Riek et al., 1996; Donne et al., 1997; Hornemann et al., 1997; Riek et al., 1997; Liu et al., 1999) (reviewed by Wüthrich and Riek, this volume). The aggregated, but noncrystalline nature of PrP-res has made it impossible to determine its high resolution structure with current technology. However, infrared (Fig. 1A), circular dichroism and x-ray diffraction analyses have shown that, in contrast to PrPC, PrPSc is predominantly β sheet with lower proportional α-helix content (Caughey et al., 1991; Safar et al., 1993; Pan et al., 1993; Nguyen et al., 1995; Caughey et al., 1998). Therefore, the transition of a portion of the α-helical and/or disordered secondary structures to β sheet is thought to be key to the conversion of PrPC to PrP-res.

FIG. 1. Strain-associated PrP-res conformations. (A) Primary and second derivative Fourier transform infrared spectra of PrP-res isolated from Syrian hamsters with the HY or DY TSE strains. Although the PrP-res in these samples are derived from the same hamster PrPC sequence, major differences are observed between the DY and HY strains in the β-sheet region of the spectrum (~1616–1640 cm–1) (Caughey et al., 1998). These differences correlate with the 1 to 2 kDa difference in the immunoblot bands after PKdigestion (Bessen and Marsh, 1994). The multiple bands represent the unglycosylated form (bottom band) and various larger glycoforms. (B) A nucleated polymerization-style model illustrating how differently conformed oligomers of PrP-res might propagate themselves. The squares represent the PK-resistant core in PrP-res (e.g., residues ~90–232 in HY PrP-res and a 1–2 kDa smaller fragment in DY PrP-res). The wavy lines designate the N-terminal domain that remains PK-sensitive in PrP-res. The heavier segment of the wavy lines in the DY PrP-res structure shows additional residues that are removed from this conformer by PK. Also shown are strain-specific, cell-free conversion reaction products of reactions between hamster 35S-PrP-sen (the largely unglycosylated GPI– construct) and HY or DY PrP-res. The difference in the size of these products after PK digestion and SDS-PAGE mimics that of the brain-derived PrP-res shown in (A). The data are consistent with the concept that different PrP-res conformations may account for the existence and individual properties of TSE strains (Caughey et al., 1998).

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Further information about the structure of PrPSc has been sought with unfolding studies (Safar et al., 1993; Oesch et al., 1994; Safar et al., 1994; Caughey et al., 1995; Kocisko et al., 1996). Initial studies showed that hamster PrPSc can be irreversibly disaggregated and unfolded with guanidine HCl (GdnHCl) or acid treatments. These studies surmised that PrPSc is fully monomerized at ~1.5 M GdnHCl and then unfolded at 2.5–3 M via a monomeric molten globule-like folding intermediate (Safar et al., 1993; Safar et al., 1994). More recent studies obtained evidence for more structural heterogeneity in PrPSc aggregates and a somewhat different unfolding/disaggregation pathway (Caughey et al., 1995; Kocisko et al., 1996) (Fig. 2). When isolated from 263K scrapieinfected hamster brain tissue without exposure to PK, at least two types of PrP molecules can be observed in PrPSc aggregates. Roughly half of the molecules appear to be fully sensitive to PK digestion and can be readily dissociated from the PK-resistant core polymers with 2.5–3 M GdnHCl or PK treatment (Caughey et al., 1995; Caughey et al., 1997). The remaining PrP-res aggregates have three distinguishable domains within the polymeric structure. The N-terminal residues 23–~89 are sensitive to PK cleavage in the absence of denaturant. Another ~3 kDa domain beginning at ~ residue 90 can be unfolded reversibly and rendered PK-sensitive by exposure to 2.5–3 M GdnHCl. Another C-terminal domain (~16 kDa in the aglycosyl structure) is the most resistant to PK as the GdnHCl concentration is increased. This domain becomes sensitive to PK at 3–4 M GdnHCl in an irreversible unfolding process that is accompanied by depolymerization and losses of both converting activity and scrapie infectivity (Caughey et al., 1997). Attempts to reverse the loss of infectivity after denaturation of PrPSc have met with variable results. McKenzie et al. (1998) reported that copper ions can assist the refolding of PrPSc after partial unfolding in GdnHCl and loss of TSE infectivity. However, other investigators have been unable to restore scrapie infectivity lost to treatments with urea, chaotropic salts, and SDS (Prusiner et al., 1993; Riesner et al., 1996; Post et al., 1998; Wille and Prusiner, 1999). For instance, Riesner et al. (1996) showed that PK-treated PrP-res fibrils (prion rods) can be disrupted with SDS to form 10-nm spherical particles with high α-helical content and no PK-resistance or infectivity. On removal of the detergent or addition of acetonitrile, high β-sheet, PK-resistant aggregates were generated but they lacked infectivity (Riesner et al., 1996; Post et al., 1998). Aggregated and PK-resistant forms of PrP, which have an amorphous ultrastructure and much reduced infectivity, have also been generated after DMSO treatment of PrPSc (Shaked et al., 1999). These and other studies (e.g., Hill et al., 1999) indicate that PK resistance and aggregation alone are not sufficient for PrP to be infectious.

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FIG. 2. Model for heterogeneity and GdnHCl-induced unfolding of PrPSc (Caughey et al., 1995; Kocisko et al., 1996). At 1.5–3 M GdnHCl, the following events happen: (1) a population of tightly bound PrP-sen molecules are stripped off ordered aggregates of PrP-res, (2) superaggregates of fibrillar or protofibrillar polymers are dissociated without depolymerization of the core polymers, and (3) a portion of each monomer within the core fibrils or protofibrils is reversibly unfolded, leaving the remaining C-terminal domain (roughly residues 121–228) folded and PK-resistant. With higher GdnHCl concentrations, the fibrils/protofibrils are depolymerized and the PrP-res molecules are irreversibly denatured. Concomitant with the latter changes are the loss of associated scrapie infectivity and PrP-res in vitro self-propagating activity (Caughey et al., 1997).

Although harsh chemical treatments can denature PrP-res polymers in vitro and reduce infectivity, it is important from a therapeutic point of view to know whether PrPSc formation is reversible under physiological conditions. One analysis has indicated that under nondenaturing conditions favorable for the conversion reaction (pH 6, 200 mM KCl, 0.6% sarkosyl), no detectable solubilization/monomerization of prewashed

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PrPSc aggregates occurred over several days (M. Callahan and B. Caughey, manuscript in preparation). Given the limits of the immunoblot detection system, the solubility of PrPSc under these conditions should be <1 nM. It will be of interest to determine whether other physiological conditions or factors such as chaperones, degradative enzymes, or pharmaceutical agents might enhance PrPSc turnover in vivo as has been observed with other types of amyloid deposits (Rydh et al., 1998). In this regard, Soto and colleagues have identified synthetic peptides dubbed “beta breakers,” which tend to destabilize PrP-res and other amyloids (Sigurdsson et al., 2000; Soto et al., 2000). On the one hand, if PrP-res is the most important neuropathogenic substance in TSE pathogenesis, then such destabilizing factors may be of therapeutic benefit. On the other hand, if smaller PrP-res aggregates are more pathogenic than larger ones, or if an intermediate or byproduct of PrP-res formation is more toxic than PrP-res itself, then destabilization of PrP-res deposits in vivo may have a detrimental rather than beneficial effect (Masel et al., 1999). B. Formation of PrP-res by Deleted Forms of PrP Studies in scrapie-infected mouse neuroblastoma cells showed that PrPSc formation occurs in cells expressing PrP lacking N-terminal residues 23–88 (Rogers et al., 1993). Recently, a 106-residue deletion mutant of PrP lacking residues 23–88 and 141–176 (∆106) was also shown to be capable of supporting infection, disease propagation, and the formation of PrPSc-like molecules when expressed in transgenic mice (Supattapone et al., 1999a). Thus, these deleted portions of the PrP molecule are not necessary for PrP’s role in this model of TSE disease at least. However, expression of PrP lacking residues 114–121 blocked PrPSc formation (Holscher et al., 1998). Thus these residues, which form a hydrophobic stretch in a region of the PrPC molecule that undergoes major conformational change in its conversion to PrP-res, appear to be critical to the conversion process. This conclusion is supported further by the inhibition of PrP-res formation by synthetic peptides containing at least some of these residues (Chabry et al., 1998; Chabry et al., 1999). C. Structural Diversity in Abnormal TSE-Associated PrP Molecules Terms such as PrP-res and PrPSc are often used as if they represent a single conformational state or entity; however, it is increasingly apparent that this is an oversimplification. TSE-associated PrP molecules can vary in resistance to proteolysis (Bessen and Marsh, 1994; Tagliavini et

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al., 1994; Bessen et al., 1995; Caughey, B. et al., 1998; Safar et al., 1998), insolubility in detergents (Somerville et al., 1989; Bessen and Marsh, 1994; Muramoto et al., 1996), secondary structure (Fig. 1A) (Caughey et al., 1998), glycoform ratios (Kascsak et al., 1986; Somerville and Ritchie, 1990; Monari et al., 1994; Collinge et al., 1996; Somerville et al., 1997; Hope et al., 1999; Rudd et al., 1999), exposure of epitopes by denaturants (Safar et al., 1998), multispectral ultraviolet fluorescence (Rubenstein et al., 1998), ultrastructure (McKinley et al., 1991; Giaccone et al., 1992; Jeffrey et al., 1994), and membrane topology (Hegde et al., 1999). Complicating matters further is the fact that PrPSc can be isolated as a complex of both partially PK-resistant and fully PK-sensitive molecules (Caughey et al., 1995)(Fig. 2). Although the PK-sensitive molecules are not required for infectivity and self-propagating activity (Caughey et al., 1997), they might play a role in neuropathogenesis (Horiuchi and Caughey, 1999). Finally, manipulations of various recombinant and mutant forms of PrP have generated numerous forms of PrP that appear to share some, but not all, of the properties of bona fide TSEassociated PrP isolated from infected tissues. A current challenge is to understand which abnormal states of PrP are relevant to various aspects of TSE transmission and/or pathogenesis. The requirements may be different for infectious versus neurotoxic forms of PrP. With these uncertainties in mind, Weissmann has proposed the use of the term PrP* to connote the putative infectious form of PrP, whatever it may be (Weissmann, 1991). III. MECHANISTIC MODELS OF PrP-RES FORMATION Before considering the experimental observations relating to the mechanism of PrP-res formation, it may be helpful to have prevalent theoretical models of PrP-res formation in mind. A wide variety of mechanisms and permutations of mechanisms have been proposed for PrP-res formation. Two that are most commonly considered are the heterodimer model (Griffith, 1967; Bolton and Bendheim, 1988; Prusiner, 1998) and the nucleation (seed)-dependent polymerization models (Griffith, 1967; Gadjusek, 1988; Jarrett and Lansbury, Jr., 1993; Lansbury, Jr. and Caughey, 1995) (Fig. 3). In its simplest rendition, the heterodimer model posits that PrP-res exists in a stable monomeric state that can bind PrPC, forming a heterodimer, and catalyze a conformational change in PrPC to form a homodimer of PrP-res. The PrP-res homodimer then splits apart to give two PrP-res monomers. Fundamental aspects of this model are that PrP-res is more stable thermody-

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FIG. 3. Mechanistic models for formation of PrP-res from PrPC. In the heterodimer and autocatalytic nucleated polyerization models, the conformational conversion of PrPC to PrP-res is rare unless catalyzed by contact with monomeric or polymeric PrP-res, respectively. In the noncatalytic nucleated polymerization model, the conformational interchange is rapid, but the PrP-res conformer is poorly populated unless stabilized by binding to a preformed, stable PrP-res polymer. See text for further explanation. The wavy lines in the PrP-sen and PrP-res structures designate the flexibly disordered portion of the structure (usually N-terminal residues 23 to ~89), which remains sensitive to PK after conversion to PrP-res.

namically than PrPC, conversion of PrPC to PrP-res is rare unless catalyzed by a preexisting PrP-res template, and the PrP-res homodimer tends to dissociate into monomers. In the nucleated polymerization type of model, oligomerization/polymerization of PrP is necessary to stabilize PrP-res sufficiently to allow its accumulation to biologically relevant levels. Spontaneous formation of nuclei or seeds of PrP-res is rare because of the weakness of monovalent interactions between PrPC molecules and/or the rarity of the conformers that polymerize. However, once formed, oligomeric or polymeric seeds are stabilized by multivalent interactions (Jarrett and Lansbury, Jr., 1993). In their various permutations, the two types of models can overlap. For instance, autocatalysis (or templating) of the conformational change in PrPC by PrP-res may be a feature of either the heterodimer or nucleated polymerization models (Fig. 3). However, the nucleated polymerization could also occur if PrPC rapidly interchanges between high α-helix and

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high β-sheet conformers, with the latter being stabilized greatly by binding to a preexisting polymer of PrP-res. In either type of model, a metastable PrPC folding intermediate might most favorably interact with PrP-res in the conversion reaction. Such an intermediate might resemble those generated from recombinant PrP-sen under acidic conditions (Swietnicki et al., 1997; Zhang et al., 1997; Hornemann and Glockshuber, 1998; Jackson et al., 1999) similar to those of endosomes or lysosomes in intact cells. A critical question from a biological standpoint is what physical state(s) of PrP-res is active in TSE disease processes in vivo. The nucleated polymerization model predicts that active PrP-res seeds could range in size from the minimum stable oligomeric nucleus to large polymers. Consistent with this prediction is the frequent observation that self-inducing converting activity (see later) and infectivity are associated with a wide size range of PrP-res aggregates, but not monomers (Prusiner et al., 1993; Hope, 1994; Caughey et al., 1995; Caughey et al., 1997). In contrast, heterodimer-type models propose that discrete monomers (or in related permutations, small discrete oligomers) are the active autocatalytic units. Aggregation of PrP-res would then be a side issue. A theoretical consideration of the kinetic consequences and likelihoods of the two models (Eigen, 1996) concluded that a strict noncooperative heterodimer model is highly unlikely. However, if a small oligomer (e.g., trimer or tetramer) served as a template in a highly cooperative autocatalytic reaction, then the model becomes more plausible. In any case, it was suggested that aggregation of PrP-res is likely to be an important “prerequisite of infection.” Mathematical modeling of the nucleated polymerization mechanism by Masel et al. (1999) revealed that systems of short polymers would grow the fastest. In familial and sporadic human TSEs, formation of an initial template or seed might be a spontaneous and stochastic event that can be potentiated by specific PrP mutations (Jarrett and Lansbury, Jr., 1993; Lansbury, Jr. and Caughey, 1995; Prusiner, 1998). In TSE diseases of infectious origin, transmission might be explained by acquisition of preformed PrP-res templates/seeds. Nonetheless, until the TSE infectious agent is fully understood, it seems prudent to remain open to the possibility that another type of agent, with or without a PrP component, is responsible for TSE transmission and the instigation of PrP-res formation in infected hosts. IV. BINDING INTERACTIONS BETWEEN PrP-SEN AND PrP-RES These and other theoretical models have long predicted that PrP-res and PrPC interact directly in the course of TSE pathogenesis. Initial empir-

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ical indications that PrP-res and PrP-sen may interact specifically came from studies in animals (Prusiner et al., 1990) and scrapie-infected tissue culture cells (Priola et al., 1994; Priola and Chesebro, 1995) (see Priola, this volume). Specific binding of PrP-res to PrP-sen as opposed to other proteins has been observed directly in lysates or culture supernatants from 35S-methionine-labeled cells expressing soluble PrP-sen lacking the GPI anchor (Horiuchi et al., 1999). Of the many labeled proteins in these preparations, only the binding of the 35S-PrP-sen (GPINEG) was detected under these conditions (pH 6.0, 1% sarkosyl) (Fig. 4). However, membrane attachment of PrPC may limit accessibility to PrP-res because binding of membrane-anchored (GPI+) PrPC to PrP-res has not been demonstrated (G. Baron and B. Caughey, unpublished data). This may be because the binding occurs via sites on PrP-sen that may be blocked by attachment to the membrane via the C-terminal GPI anchor (see later and Horiuchi et al., 1999). In addition, there may be interactions between the

FIG. 4. Selective binding of PrP-sen (GPINEG) by PrPSc. PrPSc isolated from scrapieinfected brain tissue was incubated with detergent cell lysates (postnuclear supernatants) from cells metabolically labeled with 35S-methionine. 35S-labeled proteins that were bound to the PrPSc aggregates were isolated by the protocol shown. The PrPScbound 35S-labeled proteins were compared to immunopurified 35S-PrP-sen, the total labeled proteins in the lysate, and to pellets obtained in the absence of PrPSc by SDSPAGE and phosphor-autoradiography. Adapted from (Horiuchi et al., 1999).

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PrPC polypeptide chain and membrane surfaces that block its access to PrPSc (Morillas et al., 1999). V. PrPSC-INDUCED CONVERSION OF PrP-SEN TO PrP-RES: BIOLOGICAL CONNECTIONS As also predicted by the original protein-only models for TSE agents, PrP-res not only binds PrP-sen but also induces a conversion to a partially protease-resistant state that is indistinguishable from TSE-associated PrP-res itself (Fig. 5) (Kocisko et al., 1994). The “converting activity” demonstrated in this reaction gives PrP-res at least limited self-propagating activity. Such activity would be important in its potential function as an infectious protein. So far, however, the yields of new PrP-res in these reactions has been limited to amounts that are less than or equal to the input PrP-res, so continuous self-propagation has not yet been observed (Horiuchi et al., 2000) (G.J. Raymond, L. Herrmann, and B. Caughey, unpublished observations). Nonetheless, there are numerous additional ways in which this simple cell-free conversion reaction reflects the biol-

FIG. 5. Cell-free PrP conversion reaction. The right panel is a phosphor-autoradiographic image of an SDS-PAGE gel showing only the radiolabeled hamster PrP molecules in the reaction with and without PK digestion after incubation in the presence or absence of unlabeled PrP-res. The PrP-res (PrPSc) was isolated from scrapie-infected hamster brain tissue. Details of the reaction can be found in (Kocisko et al., 1994).

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ogy of TSE disease in vivo. The PrP sequence specificity of the conversion reaction correlates with interspecies and intraspecies TSE transmissions in vivo and, as discussed later in more detail, this specificity may provide an important control point in interspecies TSE transmissions (Kocisko et al., 1995; Bossers et al., 1997; Raymond et al., 1997; Bossers et al., 2000; Raymond et al., 2000). The strain specificity of the reaction (see later) mimics PrP-res formation in vivo and provides a potential mechanism for TSE strain propagation by a protein-only mechanism (Bessen et al., 1995). The correlation between cell-free converting activity and scrapie infectivity GdnHCl denaturation studies suggests that these two parameters are related (Caughey et al., 1997). Finally, the fact that the in vitro conversion reaction has been adapted to physiologically compatible conditions suggests that it can also occur in vivo (Bessen et al., 1997; DebBurman et al., 1997; Horiuchi et al., 1999). Indeed, an in situ conversion reaction has been shown to occur within the context of intact, TSE-infected brain slices (Fig. 6) (Bessen et al., 1997). These experiments revealed that both amyloid plaque and diffuse deposits of PrP-res have the ability to induce conversion. The 35S-PrP-res product of the cell-free conversion reaction also mimics the behavior of PrPSc with regard to the reversibility of aggregation and domain structure (M. Callahan and B. Caughey, manuscript in preparation). PrP that binds to the PrPSc aggregate during the cell-free conversion reaction does not dissociate from the aggregate over the course of several days in conversion buffer lacking denaturant. Furthermore, when cell-free conversion reactions are digested with PK in the presence of 2.5 M GdnHCl, the resulting 35S-PrP-res band was ~3 kDa smaller than the band resulting from digestion without GdnHCl. This partial unfolding of the PK-resistant core of newly-converted PrP-res was at least partially reversible. These properties of the 35S-PrP-res conversion product are indistinguishable from those of PrPSc generated in scrapie-infected animals (Kocisko et al., 1996). The products of cell-free conversion reactions have not yet been shown to be infectious (Hill et al., 1999; G.J. Raymond, D. Kocisko and B. Caughey, unpublished observations). However, major technical difficulties are associated with such analyses given the limited yield of the current reactions and the inaccuracy of the infectivity bioassay. Thus, it remains to be determined whether PrP-res formation in this (or any) setting represents TSE agent propagation. Nonetheless, the fact that the conversion reactions reflect fundamental aspects of TSE biology suggest they are relevant to the molecular pathogenesis of these diseases. In addition, the cell-free conversion reactions have been used for practical purposes such as predicting host susceptibilities to TSE agents

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FIG. 6. In situ conversion reaction in brain slices on glass slides (Bessen et al., 1997). A distinct autoradiographic image is seen with scrapie-infected, but not uninfected control, brains. If the brain slices are solubilized after the conversion reaction and analyzed by SDS-PAGE/autoradiography, then PK-resistant 35S-PrP conversion products similar to those produced in cell-free conversion reactions (see Fig. 5) are observed. Higher magnification images (bottom panels) show that the pattern of in situ conversion product closely matches that of immunohistochemical staining for PrP-res in regions known to contain either amyloid plaques or diffuse apparently nonamyloid deposits.

(see later) (Raymond et al., 1997; Bossers et al., 2000; Raymond et al., 2000) and screening for anti-TSE compounds (Caughey et al., 1998; Chabry et al., 1998; Demaimay et al., 1998).

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VI. MECHANISTIC STUDIES OF THE PrP-RES-INDUCED CONVERSION REACTION The precise mechanism of the conversion reaction is not clear. However, a variety of observations warrant consideration in evaluating mechanistic models of PrP-res formation. The reaction is dependent on time and the concentrations of input PrP-res and PrP-sen (Caughey et al., 1995), although dependence on these parameters can be complex and condition-dependent (Caughey et al., 1995; Horiuchi et al., 1999). Converting activity is detected in association with PrP-res polymers but not with soluble, monomeric forms of PrP (Caughey et al., 1995; Caughey et al., 1997). The newly converted PrP-res remains associated with the PrP-res aggregates and is not released as a soluble entity (Fig. 6) (Bessen et al., 1997). The reaction is stimulated by a variety of factors that can affect protein folding including guanidine hydrochloride (Kocisko et al., 1994), urea (DebBurman et al., 1997), certain detergents (Raymond et al., 1997), chaperones (Tatzelt et al., 1996; DebBurman et al., 1997), or, in the absence of denaturants or detergents, sulfated glycans (Fig. 7) and elevated temperature (Wong et al., 2001). Kinetic analyses in the absence of denaturants provided evidence that the conversion process can be separated into two stages, first the binding of PrP-sen to PrPSc and then a slower conversion of the bound PrP-sen to PrP-res (Bessen et al., 1997; DebBurman et al., 1997; Horiuchi et al., 1999). These observations, and the formation of amyloid fibril polymers by PrP-res, are consistent with an autocatalytic or templated nucleated polymerization mechanism. However, the fact that PrP-res usually induces the conversion of only substoichiometric quantities of PrP-sen in current cell-free reactions makes the reaction less continuous than typical nucleated polymerizations of proteins or peptides. This may be a technical problem rather than a fundamental limitation of the reaction mechanism. On the other hand, since PrPres forms continuously for long periods of time in vivo, there may be important elements of the mechanism, such as cofactors or microenvironments, that remain to be elucidated (DebBurman et al., 1997; Saborio et al., 1999). A. Importance of the Disulfide Bond A subject of current debate is the importance of the PrP disulfide bond that links the second and third helices in the ordered C-terminal domain of PrPC (see the Wüthrich and Riek and Glockshuber chapters, this volume). Somerville et al. (1980), first reported that scrapie infec-

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FIG. 7. Stimulation of cell-free conversion reaction by heparan sulfate. In the absence of input (unlabeled) PrP-res there is no formation of newly formed 35S-PrP-res (lane 1). In the presence of input PrP-res, there is an 8-fold stimulation of 35S-PrP-res formation by 100 µg/ml heparan sulfate (compare lanes 2 and 3). 35S-PrP-sen representing 20% of the input radioactivity is shown in lane 4 without PK digestion. Adapted from Wong et al., (2001).

tivity is sensitive to the disulfide reducing agent 2-mercaptoethanol but only in the presence of SDS. Biochemical analyses indicated that PrPSc, like PrPC, contains an intact disulfide bond (Turk et al., 1988). Furthermore, the cell-free conversion reaction is inhibited by treatment of either PrP-sen or PrP-res with the reductant dithiothreitol (Herrmann and Caughey, 1998). These complementary lines of evidence suggest that preservation of the disulfide bond is important in infectivity and PrPSc formation. However, recent experiments by Jackson et al. (1999) showed that reduction of the disulfide bond aids in the conversion of recombinant PrP-sen to a high β sheet, PK-resistant, fibril-forming state similar to PrPSc. This so-called “β-PrP” has not been shown to be infectious, but its discovery has rekindled questions as to whether reduction of the disulfide bond (perhaps only transiently in vivo) might play a role in the PrPC to PrP-res conversion mechanism (see Glockshuber, this volume). B. Sites of Interaction Between PrP-sen and PrP-res The establishment of more physiological conditions for the cell-free conversion reaction allowed the use of anti-PrP antibodies to begin mapping the surfaces on PrP-sen that are involved in its initial binding to PrPSc (Horiuchi et al., 1999) (Fig. 8). So far only antibodies against the C-terminus of PrP (α219–232) have inhibited binding. However, removal of the α219–232 epitope from PrP-sen did not eliminate

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FIG. 8. Potential PrP-res binding sites on the NMR structure (Liu et al., 1999) of a Cterminal fragment (residues 119–231) of hamster PrP-sen (Horiuchi et al., 1999). The regular circles represent residues in this fragment that are known to be highly flexible (Riek et al., 1997; Liu et al., 1999). Other flexible unstructured residues 90–118 that were in the fragment analyzed by NMR are not shown. Two of the localized surfaces that might be blocked by binding of an antibody raised against residues 219–232 are encircled with dashed lines and marked with the approximate residue numbers. A third surface involves residues extending from 138 to ~119 along the back side of the model in the direction indicated by the arrow. The residues 119 and 120 are included because of their importance in peptide inhibition of the conversion reaction (Chabry et al., 1998). However their importance in the initial binding reaction remains to be determined. (Figure adapted from Horiuchi et al., 1999).

binding, suggesting that it is not the epitope itself that is required, but residues close to it in space that are sterically hindered by antibody binding. Surfaces that fit this description include the extended chain of residues ~119–140, helical residues ~206–223 and the loop of residues ~165 to 174 between the second β strand and the second α helix (Fig. 8) (Horiuchi et al., 1999). The importance of residues in these regions has also been implicated by the inhibition of the binding and conversion reactions by peptides corresponding to residues 119–136 (Chabry et al., 1998; Chabry et al., 1999), 166–179 and 200–223 (Horiuchi et al., 2001). Peptides corresponding to residues 109–141 and 90–145 can also form protease-resistant complexes with PrPC (Kaneko et al., 1995; Kaneko et al., 1997a).

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Amino acid substitutions in these regions have been shown to affect PrP-res formation in scrapie-infected cells (Kaneko et al., 1997b). The authors of the study proposed the existence of an unidentified factor, protein X, to explain the inhibition of PrP-res formation by point mutations in these regions. However, it also seems possible that the mutations affect the direct binding and/or conversion of PrPC to PrPSc. The ability of such mutants to inhibit PrPSc formation in trans may be due to an ability to bind wild-type PrP-res, and, without converting themselves, to block the binding and conversion of wild-type PrPC (see Section VII, C) (Priola et al., 1994; Horiuchi et al., 2000). VII. PrP-SEN/PrP-RES INTERACTIONS AND SPECIES BARRIERS The barriers to TSE transmissions between species can be strikingly high. For instance, 10 million LD50 units of 263K hamster scrapie infectivity will not kill a mouse. However, the apparent transmission of bovine spongiform encephalopathy (BSE) to humans (Bruce et al., 1997; Hill et al., 1997) and other mammalian species indicates that interspecies transmissions can occur with devastating consequences. These circumstances underscore the importance of understanding the factors that control the susceptibility of hosts to TSE agents from other species. Extensive genetic and cell culture experiments have shown that a certain degree of amino acid sequence homology between exogenous PrP-res and endogenous PrP-sen is required for the maintenance of PrP-res propagation and TSE infection (for review, see the Priola and Asante and Collinge chapters, this volume). These experiments have also shown that coexpression of incompatible PrP-sen molecules can interfere with these processes (Prusiner et al., 1990; Priola et al., 1994). A. Conversion Efficiency versus Susceptibility The sequence specificity of PrP-sen/PrP-res interactions has been investigated directly using cell-free PrP binding and conversion assays. Many “heterologous” conversion reactions have been performed using PrP-res of one species/sequence and PrP-sen of another. Profound sequence specificity is often observed (e.g., Fig. 9) (Kocisko et al., 1995; Bossers et al., 1997; Raymond et al., 1997; Chabry et al., 1999; Bossers et al., 2000; Raymond et al., 2000). In side-by-side assays with a given PrPres preparation, manyfold (5 to > 50-fold) stronger conversion efficiencies have been observed with PrP-sen molecules from highly susceptible animals versus clearly resistant species/genotypes. Intermediate effi-

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ciencies (2 to 4-fold weaker than homologous) have been observed with PrP-sen from animals that are susceptible, but apparently less so than the original host species. Based on the available information, it seems that the log of the relative intracerebral transmission titer might be roughly proportional to the relative cell-free conversion efficiency on a linear scale (Raymond et al., 2000). However, much more quantitative transmission data between various species will be required to establish the fit between these parameters. Nonetheless, the requirement for molecular compatibility between different PrP-res and PrP-sen sequences, as reflected both in vivo and in vitro, appears to be an important factor in the transmission process. B. Chronic Wasting Disease The cell-free conversion assay has been used to gain insight into the likely susceptibility of various hosts to TSE agents from different source species or genotypes (Raymond et al., 1997; Bossers et al., 2000; Raymond et al., 2000). For instance, little is known about the transmissibility of chronic wasting disease (CWD) of deer and elk to other noncervid species (Bartz et al., 1998; Miller et al., 2000). In cell-free conversion reactions, we recently found that the CWD-associated PrP-res (PrPCWD) of cervids readily induces the conversion of cervid PrP-sen molecules to the protease-resistant state in accordance with the known transmissibility of CWD between cervids (Fig. 9) (Raymond et al., 2000). In contrast, PrPCWD-induced conversions of human and bovine PrP-sen were much less efficient, and, conversion of ovine PrP-sen was intermediate. These results demonstrate a barrier at the molecular level that should limit, but not necessarily eliminate, the susceptibility of these noncervid species to CWD. C. Binding versus Conversion of Heterologous PrP Molecules Since the conversion reaction has been dissected kinetically into initial binding and conversion-to-PrP-res steps, the question arises as to which step is most important in controlling the sequence specificity of the conversion reaction (DebBurman et al., 1997; Horiuchi et al., 1999). Recent experiments using mouse and hamster PrP isoforms indicated that the binding of PrP-sen to heterologous PrPSc can occur much more efficiently than the subsequent conversion to PrP-res (Horiuchi et al., 2000). This suggests that the species-specificity of PrP-res formation is determined more by the conversion step than the initial binding step. The binding of heterologous, nonconverting PrP-sen molecules to

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PrPSc does not block the binding of homologous PrP-sen, but does interfere with its conversion. This interference effect could explain reductions in the rates of PrP-res formation and/or TSE pathogenesis in hosts that coexpress heterologous PrPC molecules (Prusiner et al., 1990; Palmer et al., 1991; Priola et al., 1994). As noted previously, the binding and conversion steps may occur at a single site (Fig. 10), or there may be two types of PrP-sen binding sites on PrP-res, one that is “conversion-competent” (say, at the ends of growing fibrils) and another that is not (e.g., along the sides of fibrils as depicted in Fig. 2) (Caughey et al., 1995; Caughey et al., 1997). The initial binding of PrP-sen to PrP-res at either type of site may occur in a manner that is less amino acid sequence-specific than the additional steps required at conversion-competent sites for the conversion of PrPsen to the PK-resistant state (Horiuchi et al., 2000). Studies using mouse/hamster chimeric PrP have shown that the central part of the PrP-sen molecule, including three amino acid substitutions at mouse/hamster residues 138/139, 154/155, and 169/170, is important in the conversion of PrP-sen to PrP-res (Scott et al., 1992; Priola et al., 1994; Kocisko et al., 1995; Priola and Chesebro, 1995). Thus, it is possible that critical interactions in the vicinity of these residues on PrP-sen and/or PrPSc occur as part of the conversion step. VIII. TSE STRAINS AND PrP-SEN/PrP-RES INTERACTIONS TSE strains can be discriminated within a single host species or PrP genotype by reproducible differences in incubation period, clinical signs, vacuolar brain pathology, and PrP-res deposition (for review, see Bruce and Fraser 1991.) The existence of TSE strains independent of the host PrP genotype presents a major challenge to the protein-only

FIG. 9. Efficiencies of conversion reactions between homologous and heterologous isoforms of PrP. Conversion efficiencies of [35S]-PrP-sen proteins in cell-free reactions with equivalent amounts of different PrP-res molecules. The results show the mean percentage of the input 24–27 kDa [35S]-PrP-sen converted to the 17–20 kDa PK- resistant [35S]-PrP bands. PrPCJD-M/M and PrPCJD-V/V designate human PrP-res derived from the brains of CJD patients homozygous for methionine or valine at residue 129, respectively. Within the groups of values for each type of PrP-res, the statistical significance of the difference of the means relative to the maximum mean percent conversion of PrPsen of the same species (boxed) was assessed with a one way ANOVA with Dunnett’s multiple comparison test: *, p < 0.05; **, p < 0.01. The “norm mean” column shows means within each PrP-res group normalized to the homologous conversion (boxed). Adapted from Raymond et al., 2000.

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FIG. 10. Nucleated polymerization-based model for binding, conversion (top), and interference (bottom) phenomena that is consistent with previous experimental observations (DebBurman et al., 1997; Horiuchi et al., 1999; Horiuchi et al., 2000). This model assumes a single site for both binding and conversion, but a two-site model is described elsewhere (Horiuchi et al., 2000). Rapid binding of PrP-sen (open triangles) is followed by a slower wave of conformational conversion of bound PrP-sen molecules to PrP-res (squares). The inclusion of nonconvertible PrP-sen (black triangles) among convertible PrP-sen molecules interferes with propagation of the conversion through the bound PrP-sen molecules without preventing binding of the homologous PrP-sen (bottom) (Horiuchi et al., 2000). PK designates a proteinase K digestion step wherein PrP-sen is completely digested and the N-terminal octapeptide repeat domain (residues 23-~90, the wavy lines in the PrP-sen and PrP-res structures) are removed from PrP-res. (Adapted from Horiuchi et al., 2000).

hypothesis for the TSE agent. However, there is evidence to support the hypothesis that the mechanism for TSE strain propagation is determined by the self-propagation of PrP-res molecules that differ in conformation, glycoform ratios, polymeric states, and/or ligand associations (Bessen and Marsh, 1994; Bessen et al., 1995; Collinge et al., 1996; Telling et al., 1996; Somerville et al., 1997; Caughey et al., 1998; Rubenstein et al., 1998; Safar et al., 1998). Kascsak and colleagues first observed that PrP-res associated with various murine scrapie strains differ in the relative proportions of PrP-res glycoforms (i.e., PrP molecules with 0, 1 or 2 N-linked glycans) (Kascsak

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et al., 1985; Kascsak et al., 1991). The possibility of multiple conformations of PrP-res was suggested by observations that PrP-res associated with different strains of TSEs are cleaved at different N-terminal sites by PK. This was first documented with the HY and DY strains of transmissible mink encephalopathy (TME) passaged in hamsters (Bessen and Marsh, 1992; Bessen and Marsh, 1994; Bartz et al., 2000). After treatment with PK, the PrP-res from HY-infected hamsters had a ~2-kDa larger molecular mass than PrP-res from DY-infected hamsters in SDSPAGE analysis. N-terminal sequencing revealed that PK cleaved additional residues from the N-terminus of DY PrP-res compared to HY PrP-res. Since the DY and HY PrP-res molecules are derived from the same Syrian hamster PrP-sen precursor, it was concluded that the observed difference in cleavage by PK was due to strain-dependent differences in conformation and/or ligand binding. Further evidence for conformational differences has come from FTIR spectroscopy, which indicated that HY and DY PrP-res differ dramatically in the beta sheet region (Caughey et al., 1998) (Fig. 1A). More recently, related, but more subtle differences have been observed among spectra of various strains of mouse PrPSc (B. Caughey and G. Raymond, unpublished data). Collinge and colleagues have shown that metal ion binding to PrP-res can influence the site at which it is cleaved by PK (Wadsworth et al., 1999). Evidence for multiple strain differences in PrPSc conformation has also been obtained with multispectral ultraviolet fluorescence spectroscopy (Rubenstein et al., 1998) and by comparing antibody epitope exposure caused by denaturant treatments (Safar et al., 1998). Evidence that the two strain-associated conformations of hamster PrP-res could propagate themselves from the same hamster PrP-sen precursor was obtained using a cell-free conversion reaction (Fig. 1B) (Bessen et al., 1995). In these studies, HY and DY PrP-res were each incubated with hamster PrP-sen, and the PrP-sen was converted to PKresistant PrP products with the same ~1–2 kDa difference in molecular mass that distinguishes the PK-treated HY and DY PrP-res molecules. This finding suggested that the strain-specific forms of PrP-res are faithfully propagated through direct PrP-sen-PrP-res interactions both in vitro and in vivo as had been proposed earlier (Bolton and Bendheim, 1988; Prusiner, 1991; Bessen and Marsh, 1994). Strain-dependent differences in PrP-res have been observed with other TSEs, and these differences now serve as a biochemical aid in TSE strain typing (see Asante and Collinge, this volume) (Monari et al., 1994; Collinge et al., 1996; Telling et al., 1996; Parchi et al., 1997; Somerville et al., 1997; Bartz et al., 2000). Indeed, this type of analysis

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has provided corroborative evidence connecting the bovine spongiform encephalopathy strain of TSE in cattle to the variant CreutzfeldtJakob disease (CJD) in humans (Hill et al., 1997; Somerville et al., 1997a). Using a similar type of analysis, the transmission of different strains of human CJD into mice leads to the formation of mouse PrP-res with different PK cleavage sites (Telling et al., 1996). These observations show that the strain-specific conformations can be propagated between, as well as within, species. IX. PrP PERTURBATIONS AND TSE INFECTIVITY Although PrP-res is associated with TSE infectivity and has, at the biochemical level at least, some limited self-propagating activity, important questions remain about the PrP-only prion model for the TSE agent. For instance, why has no one been able to demonstrate the model’s most basic prediction, that is, that PrP-sen alone can be induced to convert to an infectious agent that causes TSE disease when inoculated into animals? Inoculation of a synthetic PrP peptide has induced a TSE-like disease in transgenic mice expressing a familial TSE associated mutant PrP, but the serial transmissibility of this disease remains to be determined (Kaneko et al., 2000). Why are ~100,000 PrP-res molecules present per infectious unit? Why is PrP-res not always found in infectious tissues (Lasmezas et al., 1997)? Why are nucleic acids (Akowitz et al., 1990), polysaccharide (Appel et al., 1999), and lipids (Klein et al., 1998) common “contaminants” of PrP-res preparations and what role might they play in infectivity? Why has cell-free conversion of PrP-sen by PrPres been limited to substoichiometric yields? Since these unanswered questions may have purely technical explanations, they do not negate the potential for PrP-res to act as a self-propagating prion. However, they indicate that fundamental mysteries remain about molecular mechanisms of TSE disease. X. PrP-SEN/PrP-RES INTERACTIONS AND THE SEARCH FOR ANTI-TSE DRUGS The development of effective treatments for TSE diseases remains problematic. A number of compounds have been identified that can prolong the lives of scrapie-infected rodents if administered before or near the time of infection. These compounds include various polyanions (e.g., pentosan polysulfate (Kimberlin and Walker, 1986; Lado-

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gana et al., 1992), dextran sulfate (Ehlers and Diringer, 1984; Farquhar and Dickinson, 1986; Kimberlin and Walker, 1986), HPA23 (Kimberlin and Walker, 1983), Congo red (Caughey et al., 1993; Ingrosso et al., 1995), porphyrins, and phthalocyanines (Priola et al., 2000). Unfortunately, these compounds do not appear to cross the blood-brain barrier and have no effect if administered after the infection has entered the central nervous system. Polyene antibiotics (e.g., amphotericin B and MS8209) have been shown to have beneficial effects much later in the course of infection, but still have little or no effect after the appearance of clinical disease (Demaimay et al., 1994; Adjou et al., 1995; Demaimay et al., 1997; Demaimay et al., 1999). Furthermore, the amphotericins tend to be too toxic for application to TSE diseases in humans. Other compounds such as the “beta breaker peptides” (Soto et al., 2000) and anthracycline (Tagliavini et al., 1998) can neutralize infectivity if incubated directly with the inoculum before inoculation, but are not known to be effective if given to the host postinoculation. Thus, there is still a dire need for effective anti-TSE drugs. Most of the antiscrapie agents listed previously have been shown to be inhibitors of PrP-res formation in scrapie-infected neuroblastoma cells (Caughey and Race, 1992; Caughey et al., 1993; Caughey and Raymond, 1993; Caughey et al., 1998). Indeed, Congo red and the porphyrins and phthalocyanines were first identified as potential antiscrapie drugs in scrapie-infected tissue culture cells (Caughey and Race, 1992; Caughey and Raymond, 1993; Demaimay et al., 1998; Rudyk et al., 2000; Demaimay et al., 2000). Other classes of compounds such as synthetic PrP peptide fragments (Chabry et al., 1998; Chabry et al., 1999), lysosomotropic amines (e.g., quinacrine) (Doh-ura et al., 2000), cysteine protease inhibitors (Doh-ura et al., 2000), branched polyamines (Supattapone et al., 1999b), cationic lipopolyamines (Winklhofer and Tatzelt, 2000) and mimics of dominant negative inhibitors of scrapie replication (Perrier et al., 2000) have been shown to have antiscrapie activity in this system, but have not been tested in animals. The track record so far indicates that inhibitors of PrP-res formation in vitro often have anti-TSE activity in vivo. Testing in animals is ultimately essential for characterization of potential anti-TSE drugs; however, it is slow, cumbersome, and expensive. Thus, the development of higher throughput screens for inhibitors of PrP-res formation or the PrP-sen-PrP-res interactions that are required for PrP-res formation should aid in the search for new potential drugs. Of particular interest would be inhibitors that can cross the blood-brain barrier to potentiate therapeutic activity after TSE infections have spread to the central nervous system.

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XI. CONCLUSIONS The properties of PrP-res have foiled attempts to determine its high resolution structure. Nonetheless, low resolution analyses have shown that PrP-res retains significant α-helical content but has much higher βsheet content than PrPC. Biochemical analyses indicate that within its ordered aggregate, PrPSc molecules have at least three distinguishable folding domains. The ~6-kDa N-terminal domain remains flexibly disordered and PK-sensitive. A ~3-kDa middle domain (approx. residues 90–120) is ordinarily PK-resistant but can be unfolded reversibly in 2.5 M GdnHCl. The C-terminal domain containing the N-linked glycans, α helices, and the GPI anchor is the most stable to GdnHCl-induced denaturation. The ordered PrPSc aggregates can be either ultrastructurally indistinct or in the form of amyloid fibrils and plaques. PrP-res aggregates can selectively bind PrP-sen via a localized site close in space to the C-terminus. Under physiologically compatible conditions, PrPres can also induce the conversion of PrP-sen to a protease-resistant state reminiscent of PrPSc. This conversion reaction is highly speciesand strain- specific, as is PrP-res formation in vivo. Heterologous PrPsen molecules from one species can bind to PrP-res of another and interfere with the conversion of homologous PrP-sen molecules. It is not known whether the cell-free conversion reaction, or any other in vitro manipulation of PrP-sen, generates infectivity. Thus, the full nature of the agent remains unclear. Nonetheless, the accumulation of aberrant forms of PrP, such as PrP-res, seems to be critical in the TSE pathogenesis. Inhibitors of PrP-res formation can prolong the lives of scrapie-infected animals if given near the time of infection, but no acceptable drugs have been identified that are effective once clinical signs of TSE disease have appeared. REFERENCES Adjou, K.T., Demaimay, R., Lasmezas, C., Deslys, J.-P., Seman, M., and Dormont, D. (1995). Antimicrob. Agents Chemother. 39, 2810–2812. Akowitz, A., Sklaviadis, T., Manuelidis, E.E., and Manuelidis, L. (1990). Microb. Pathog. 9, 33–45. Appel, T.R., Dumpitak, C., Matthiesen, U., and Riesner, D. (1999). Biol. Chem. 380, 1295–1306. Bartz, J.C., Marsh, R.F., McKenzie, D.I., and Aiken, J.M. (1998). Virology 251, 297–301. Bartz, J.C., Bessen, R.A., McKenzie, D., Marsh, R.F., and Aiken, J.M. (2000). J. Virol. 74, 5542–5547. Bessen, R.A., Kocisko, D.A., Raymond, G.J., Nandan, S., Lansbury, P.T., Jr., and Caughey, B. (1995). Nature 375, 698–700.

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STUDIES ON PEPTIDE FRAGMENTS OF PRION PROTEINS BY FABRIZIO TAGLIAVINI, GIANLUIGI FORLONI,* PASQUALINA D’URSI,* ORSO BUGIANI, AND MARIO SALMONA* Istituto Nazionale Neurologico Carlo Besta and *Istituto di Ricerche Farmacologiche Mario Negri, Milano, Italy

I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. The Amyloid Peptides of Gerstmann-Sträussler-Scheinker Disease . . . . . . . . . III. Unraveling the Conformational Conversion of PrPC to PrPSc Using Synthetic Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The Flexible N-terminal Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. The Globular Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Unraveling the Pathogenesis of Prion Diseases Using Synthetic Peptides . . . A. Interaction of PrP Peptides with Cell Membranes . . . . . . . . . . . . . . . . . . . . B. Effects of PrP Peptides on Neuronal Cultures . . . . . . . . . . . . . . . . . . . . . . . C. Effects of PrP Peptides on Astroglial and Microglial Cultures . . . . . . . . . . V. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. INTRODUCTION The prion diseases are neurodegenerative disorders of humans and animals that are sporadic or inherited in origin and can be transmitted (Prusiner, 1991). Despite remarkable differences in phenotypic expression, these disorders share a similar pathogenic mechanism, that is, a posttranslational modification of the prion protein from a normal cellular isoform (PrPC) to disease-specific species (PrPSc). Nuclear magnetic resonance (NMR) studies of recombinant murine, hamster, bovine, and human PrP indicate that the normal protein is composed of two structurally distinct moieties: an extended N-terminal segment (residues 23–125) with features of a flexibly disordered polypeptide chain, and a well-defined globular domain (residues 126–231) with three α helices and two-stranded antiparallel β sheet (Fig. 1) (Riek et al., 1996; Riek et al., 1997; Donne et al., 1997; Lopez Garcia et al., 2000; Zahn et al., 2000). The PrPC>PrPSc transition involves a profound conformational change with decrease in α-helical secondary structure (~40% > 30%) and striking increase in β-sheet content (~3%>40%) (Caughey et al., 1991; Pan et al., 1993; Safar et al., 1993). This rearrangement is accompanied by the acquisition of abnormal physicochemical properties, including insolubility in nondenaturing detergents and 171 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

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FIG. 1. Schematic representation of PrPC, of amyloid peptides isolated from GSS brains, and a partially deleted PrP sequence that supports prion propagation. The top diagram illustrates the polypeptide chain of mature human PrP with the ocatapeptide repeat region spanning residues 51–90 and the common M/V polymorphism at codon 129. The normal protein is composed of two structurally distinct moieties, that is, a flexible N-terminal segment (residues 23–125) and a globular domain (residues 126–231) with three α helices (black areas) and two stranded antiparallel β sheets (arrows). The second and third diagrams correspond to amyloid peptides isolated from GSS patients carrying A117V, F198S, or Q217R mutation in coupling phase with valine at codon 129. The major N- and C-terminal signals obtained from protein sequence analysis and the amino acid residue at position 129 are indicated. The amyloid peptides originated from mutant molecules, because only valine was found at position 129, although the patients were M/V heterozygous at that codon. The last diagram illustrates a redacted version of PrP lacking residues 23–88 and 141–176, which is sufficient to support prion disease.

partial resistance to proteinase K digestion (Prusiner, 1991). In the presence of detergents, the protease-resistant core of PrPSc, which is Nterminally truncated near residue 90, assembles into insoluble fibrillary structures that exhibit the tinctorial and ultrastructural properties of amyloid (Prusiner et al., 1983). Thus, prion diseases share a basic molecular mechanism with amyloid diseases in that they both involve the conversion of a normally soluble form of a protein into an insoluble quaternary structure having an extensive β-sheet conformation. PrP amyloid formation occurs consistently and to the highest degree in two human disorders, (i.e., Gerstmann-Sträussler-Scheinker disease [GSS] and PrP-cerebral amyloid angiopathy [PrP-CAA]) (Ghetti et al., 1996a). In the former, amyloid deposits are essentially parenchymal, whereas in the latter, they are predominantly vascular. Deposition of PrP amyloid in the neuropil is also a consistent feature of the new vari-

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ant of Creutzfeldt-Jakob disease (CJD), which seems to be causally linked to bovine spongiform encephalopathy (Will et al., 1996; Bruce et al., 1997). In all these conditions, amyloid fibrils are associated with PrP aggregates that do not exhibit the tinctorial, optical, and ultrastructural properties of amyloid. The latter are the only or the prevalent form of PrP deposits in sporadic and familial CJD, fatal familial insomnia (FFI), and natural prion disease of animals (e.g., scrapie of sheep and goat and spongiform encephalopathy of cattle). Whether these nonfibrillar deposits possess an underlying regular molecular structure is unknown. Co-occurrence of nonfibrillar and fibrillar amyloid deposits has been observed in many other conditions such as amyloid-β (Aβ) and immunoglobulin light chain aggregation disorders (Bugiani et al., 1989; Kaplan et al., 1997; Miravalle et al., 2000). Since at least in some instances protein aggregation in vivo is based on self-association of partially folded intermediates, different sequential or divergent folding intermediates might be the building blocks for nonfibrillar and fibrillar PrP aggregates. The existence of different abnormal conformers of PrPSc has been inferred by the observation that distinct protease-resistant fragments of PrPSc are found in vivo and can be generated in vitro by limited proteolysis with proteinase K (Bessen and Marsh, 1992; Bessen and Marsh, 1994; Parchi et al., 1996; Piccardo et al., 1996; Piccardo et al., 1998). It is noteworthy that these PrPSc types and fragments thereof are linked to distinct neuropathological profiles. In particular, amyloid formation is consistently present in conditions, such as GSS and PrP-CAA, in which PrPSc undergoes a specific proteolytic processing that originates low-molecular-weight, N- and C-terminal truncated peptides (Tagliavini et al., 1991; Tagliavini et al., 1994; Piccardo et al., 1996; Piccardo et al., 1998). On the other hand, nonfibrillar protein aggregates, which are the prevalent form of PrPSc deposition in CJD, FFI and natural prion diseases of animals, are associated with fulllength PrPSc and N-terminal truncated peptides. The recognition of the protein domains involved in the conformational change of PrPC into PrPSc and the structure of PrPSc conformers has been limited by the insolubility of these molecules. To gain insights into this issue, physicochemical studies have been carried out using synthetic or recombinant fragments of PrP corresponding to regions of putative α-helical structure as deduced by structural predictions. An alternative approach was to design synthetic peptides homologous to the PrP region essential for amyloid fibril formation, as amyloid proteins have a high content of β-sheet secondary structure—the central feature that distinguishes PrPSc from PrPC—and are often small fragments of a precursor protein.

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II. THE AMYLOID PEPTIDES IN GERSTMANN-STRÄUSSLER-SCHEINKER DISEASE GSS disease is an autosomal dominant disorder linked to variant PRNP genotypes resulting from the combination of a pathogenic mutation (P102L, P105L, A117V, F198S, D202N, Q212P, Q217R, 192 bp insertion corresponding to 8 extra copies of the octapeptide repeat) with a common polymorphism at codon 129 (M/V) (Young et al., 1999). In most instances, specific GSS genotypes are associated with distinct clinicopathological phenotypes, a circumstance that is particularly relevant for unraveling the ground of phenotypic variability (Ghetti et al., 1996a). The biochemical composition of PrP amyloid was first determined in patients of the Indiana kindred of GSS, carrying a F>S substitution at PrP residue 198 in coupling phase with V129. Protein fractions extracted from amyloid cores isolated from cerebral cortex contained two major peptides of ~11 and ~7 kDa, spanning residues ~58–150 and ~81–150 of PrP, respectively (Fig. 1) (Tagliavini et al., 1991; Tagliavini et al., 1994). To verify whether these peptides originated from mutant PrP or from both mutant and wild-type molecules, M/V heterozygotes at codon 129 were selected for study, and V129 was used as a marker of the mutant allele. Amino acid sequencing and mass spectrometry showed that the purified amyloid fractions contained only peptides with V129, suggesting that only mutant PrP was involved in amyloid formation (Tagliavini et al., 1994). The analysis was subsequently extended to GSS patients with other PRNP mutations (i.e., A117V and Q217R). In all instances the major amyloid subunit was a ~7 kDa N- and C-terminal truncated fragment of PrP of similar size and sequence, which was derived from the mutant allele (Tagliavini et al., 1994; Tagliavini et al., 2001); in GSS A117V, the amyloid protein contained the mutant V117. The finding that the GSS amyloid protein is an internal fragment of PrP was verified by immunostaining brain sections with antisera to synthetic peptides homologous to different PrP domains (Giaccone et al., 1992). The amyloid cores were strongly immunoreactive with antisera to the mid-region of the molecule, and the periphery of the cores was immunostained by antibodies to N- or C-terminal domains. In addition, the antisera to PrP labeled large areas of the neuropil that did not show the tinctorial and optical properties of amyloid, suggesting that amyloid deposition in GSS is accompanied by accumulation of PrP peptides, which are not assembled into amyloid fibrils. Evidence suggests that N- and C-terminal cleavage of abnormal PrP isoforms that generate amyloid peptides occurs before, rather than after, fibril formation. Western blot analysis of total brain extracts from

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F198S patients has revealed three major protease-resistant PrP fragments of 27–29, 18–19, and 8 kDa. The 18–19 and 8 kDa peptides are Nand C-terminal truncated, as deduced by their antigenic profile, likely representing amyloid protein precursors (Piccardo et al., 1996). These low-molecular-weight PrP fragments are also present in areas without amyloid deposits, suggesting that brain regional factors play a key role in amyloidogenesis. Identical PrP peptides are found in patients with D202N and Q217R mutations, whose clinicopathological profile is similar to that of GSS F198S (Ghetti et al., 1996a; Lievens et al., 1998; Piccardo et al., 1998). N- and C-terminal truncated fragments are also a consistent feature of GSS P102L, A117V, and Q212P, although they can be distinguished from those of GSS F198S, D202N, and Q217R by slight differences in migration pattern (Parchi et al., 1998; Piccardo et al., 1998). This suggests structural variations of the relevant precursor protein, which could account for the differences in phenotypic expression observed in these conditions. The identification of the minimal PrP segment involved in amyloid formation in GSS (i.e., residues ~81 to ~150) along with the observations that truncation of the N-terminus (residues 23–88) and deletion of residues 141–176 do not prevent the conversion of PrPC to PrPSc (Muramoto et al., 1996; Supattapone et al., 1999) suggest that, at least in some instances, the region comprising residues ~90–140 plays a major role in the conformational change of PrP. This hypothesis is supported by in vitro studies with synthetic peptides. III. UNRAVELING THE CONFORMATIONAL CONVERSION OF PrPC TO PrPSC USING SYNTHETIC PEPTIDES Initial studies with synthetic PrP peptides focused on the identification of protein domains that could be involved in the conformational conversion of PrPC into disease-associated species. To test the hypothesis that this conversion involves the transition of α helices into β sheet, Gasset et al. (1992) examined the physicochemical properties of peptides homologous to residues 109–122, 129–141, 178–191, and 202–218 of Syrian hamster PrP, which were predicted to correspond to α-helical regions (designated H1, H2, H3, and H4, respectively) by computational studies. At variance with the theoretical predictions, peptides H1, H3, and H4 exhibited extensive β-sheet structure in the solid state and in aqueous solution and assembled into fibrils showing ultrastuctural and tinctorial properties of amyloid, as described in detail later. Multidimensional heteronuclear NMR has revealed that the prediction of the two C-terminal

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helices was largely correct, because the recombinant Syrian hamster PrP contains three helical domains spanning residues 144–156, 172–193, and 200–227. Conversely, the predicted H1 region, which showed the highest propensity to adopt extended β-sheet conformation when synthesized as peptide, does not form part of any stable secondary or tertiary fold in the entire protein and is highly flexible (Donne et al., 1997). This is consistent with the observation that PrPC to PrPSc transition is accompanied by remarkable increase in β-sheet structure and only slight decrease in αhelical content, suggesting that the conformational rearrangement occurs primarily in unstructured regions. A different strategy for identifying PrP regions involved in the conformational conversion of PrPC into PrPSc was based on physicochemical characterization of synthetic peptides homologous to consecutive segments of the GSS amyloid protein. This approach issued from the observation that the amyloid subunit was a relatively small fragment of PrP (residues ~81 to ~150) with high intrinsic ability to form insoluble quaternary structures with extensive β-sheet conformation. Furthermore, such peptides could allow in vitro experiments to investigate the molecular basis of neuronal degeneration and glial activation associated with extracellular deposition of PrP amyloid. These studies showed that the sequence spanning residues 106–147 is central to amyloid fibril formation (Tagliavini et al., 1993). In particular, a peptide homologous to residues 106–126 of human PrP (PrP106–126) exhibited high propensity to adopt stable β-sheet secondary structure and to assemble into straight, unbranched, ~8-nm-diameter filaments, ultrastructurally similar to those observed in GSS patients (Fig. 2). The fibrillary assemblies were partially resistant to protease digestion, displayed tinctorial properties of in situ amyloid (i.e, bi-refringence under polarized light after Congo red staining and yellow fluorescence after thioflavine S treatment) and showed x-ray diffraction patterns consistent with that of native amyloid fibrils (Fig. 2) (Selvaggini et al., 1993; Tagliavini et al., 1993). Most interestingly, PrP106–126 was toxic to neurons in culture, whereas it induced activation of glial cells (Forloni et al., 1993; Forloni et al., 1994). These data suggested that the distal part of the flexible N-terminal domain of PrP may be central to both the process of PrPC>PrPSc transition and the pathogenic properties of abnormal PrP isoforms. These observations and the finding that amino acid substitutions associated with genetic forms of CJD, such as D178N and E200K, increase the fibrillogenic properties of PrP fragments (Goldfarb et al., 1993) promoted a variety of studies with synthetic peptides homologous to segments of most PrP regions (Fig. 3, see color insert). The following sections review structural studies with peptides derived from the

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FIG. 2. Structural features of PrP106–126 assemblies. (A) Electron micrograph of fibrils generated by the peptide in physiological conditions. Magnification bar = 100 nm. (B) X-ray diffraction pattern of PrP106–126, with reflections corresponding to H-bonding between antiparallel β sheets (arrow).

flexible N-terminal domain and the globular C-terminal domain of PrP, and the subsequent chapter focuses on the biological properties of PrP peptides in vitro. A. The Flexible N-terminal Domain 1. The Octapeptide Repeat Region (Residues 51–90) This region, which contains a series of unusual glycine-rich repeats without significant homology to other known protein sequences, is among the most conserved parts of PrP in mammals, and analogous sequences are present in other prion proteins (e.g., hexameric repeats in chicken PrP) (Harris et al., 1991; Gabriel et al., 1992). In humans, the repeat region is composed of one nonapeptide (termed R1) followed by four octapeptides (termed R2, R2, R3, R4) that have the same amino acid sequence (PHGGGWGQ) but can be distinguished by variations in the DNA sequence (Fig. 3A). Deletion of a single octarepeat (R2 or R3) is present in about 1% of the population and does not have deleterious consequences. By contrast, the insertion of one, two, four, five, six, seven, eight, or nine extra octapeptides is associated with inherited prion disease (for review see Young et al., 1999). Based on the observation that the octapeptide repeats share the sequence PHG with the histidine-rich glycoprotein, which is thought to be involved in plasma copper transport, Hornshaw et al. (1995a) hypoth-

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esized that this region could be a metal-binding domain. To test this hypothesis, they generated synthetic peptides containing three or four copies of the octarepeat and tested their ability to bind metal ions using mass spectrometry and size exclusion chromatography. They found that the octapeptide repeat sequence preferentially binds copper over other divalent ions such as zinc, manganese, cobalt, nickel, iron, calcium, and magnesium. This finding is consistent with the observation that fulllength PrPC, but not N-terminal-truncated PrPC, can be purified by Cu2+ chelate chromatography (Pan et al., 1992). However, whether PrP has a significant role in copper binding and cuproenzyme activity—e.g., Cu/Zn superoxide dismutase—in the brain is unclear (Brown et al., 1997; Waggoner et al., 2000). Subsequently, Hornshaw et al. (1995b) investigated the secondary structure of a synthetic peptide containing four tandem repeats (PrP residues 60–91). Circular dichrosim (CD) spectroscopy revealed a random coil conformation that was independent from absence or presence of Cu2+. Miura et al. (1996) did not detect any regular structure in a single octapeptide unit in the absence of divalent metal ions using Raman spectroscopy. However, Cu2+ binding to the HGGG motif of the octapeptide induced a conformational transition in the adjacent C-terminal portion from an irregular loop to an α helix. The analysis of a longer segment comprising an octapeptide and the succeeding 12 amino acids (PrP residues 84–103) showed that the α-helical structure nucleated by Cu2+ binding in the octapeptide extended over the C-terminal sequence. Stöckel et al. (1998) investigated the metal-binding properties of a nearly full-length recombinant hamster PrP (residues 29–231). After purification, the polypeptide chain was refolded into a predominantly α-helical structure resembling that of PrPC isolated from hamster brain, as deduced by near-UV CD spectroscopy. On heating, the recombinant protein could be converted into a β-sheet-rich form. CD and tryptophan fluorescence spectroscopy revealed that the addition of Cu2+ induced a structural change in hamster PrP29–231. Furthermore, the copper-complexed PrP was more prone to shift from a soluble α-helical structure to a β-sheet aggregate on heat denaturation. The study also confirmed that the metal-binding sites of PrP are highly specific for Cu2+, as no change in tryptophan fluorescence emission was observed with other divalent cations, and pointed to a role for histidine as a chelating ligand on the basis of pH titration of copper binding. Equilibrium dialysis experiments indicated a binding stoichiometry of two copper molecules per PrP molecule at physiologically relevant concentrations, with a dissociation constant of 14 µM. This finding was at vari-

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ance with previous data suggesting 5.6 Cu2+ binding sites per PrP molecule (Brown et al., 1997). To further investigate the site and mode of binding of Cu2+ to PrP, Viles et al. (1999) analyzed a series of peptides containing two, three, and four octarepeats using various spectroscopic techniques including CD, NMR, absorption spectroscopy, and electron spin resonance spectroscopy. They found that two octapeptides bind a single Cu2+ ion with a Kd of 6 µM, whereas four octapeptides cooperatively bind four Cu2+ ions. In the absence of Cu2+, all peptides were unstructured in solution; however, a distinctive structuring consistent with turns and structured loops, without indication of α helix or β sheet, was observed on addition of Cu2+. The authors suggested that the discrepancies between these and previous data could be due to experimental conditions, in particular pH below 7.4, which is the optimum for copper binding, and use of Tris buffer, which competes for copper with the octarepeat peptides at pH 7.4. Based on spectroscopic data, Viles et al. (1999) proposed a model in which the four histidines in successive octarepeats form a bridged copper complex, with the Nε2 and Nδ1 imidazole nitrogens from each histidine coordinating two adjacent copper ions. This model would imply that, although the octarepeat region is unstructured in the absence of Cu2+, it could be constrained by the four copper-coordinating histidines into a rather compact structure at physiological Cu2+ concentrations. Based on Raman and absorption spectroscopic analysis of peptides containing one, two, and four octarepeats, Miura et al. (1999) suggested a different copper-binding mode. In this model, at neutral or basic pH, each octapeptide unit binds a Cu2+ ion via the Nπ atom of the histidine side chain and two deprotonated amide nitrogens in the adjacent triglycine segment. However, at weakly acidic pH, the Cu2+-amide– bonds dissociate and only the histidine residues provide metal coordination sites, resulting in decreased affinity of the protein to Cu2+. Because copper stimulates endocytosis of PrPC from the cell surface (Pauly and Harris, 1998), the pH-dependent change of binding mode may play a role in internalizing Cu2+ ions from extracellular to endocytic compartments. Furthermore, this change may allow formation of His-Cu-His bridges between different peptide chains, resulting in peptide aggregation. The octapeptide repeat region is not required for the conversion of PrPC into PrPSc and propagation of the disease process. Nevertheless, extra-repeat insertional mutations in one PRNP allele trigger disease. It is conceivable that disorder of copper binding in these extended protein species might feature in the formation of altered PrP isoforms and/or PrP aggregation. In this regard, it is noteworthy that PrPSc is

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extracted from brain tissue of subjects with prion disease in a metal-ionoccupied form and can undergo conformational change as a result of binding metal ions (Wadsworth et al., 1999). 2. The Distal Part of the N-terminal Domain (Residues 90–125) This region corresponds to the N-terminal part of the protease-resistant core of PrPSc (Prusiner et al., 1984). Protection against proteinase K digestion suggests that it undergoes a conspicuous structural rearrangement in PrPC to PrPSc conversion. On the other hand, mutant PrP lacking parts of this sequence (i.e., residues 95–107, 108–121, or 114–121) is not converted into a protease-resistant form in scrapieinfected mouse neuroblastoma cells (Muramoto et al., 1996; Hölscher et al., 1998). Further, overexpression of the deletion mutant PrP∆114–121 inhibits in a trans-dominant fashion the accumulation of endogenous PrPSc in these cells (Hölscher et al., 1998). Structural analysis has shown that peptides derived from the C-terminal half of this region, which is the most conserved PrP sequence across all species, are able to adopt different conformations in distinct environment, although intrinsically prefer a β-sheet structure (Fig. 3B). The first peptide to be investigated corresponded to residues 109–122 of Syrian hamster PrP (Gasset et al., 1992). Fourier transform IR (FTIR) spectra of hydrated peptide films were marked by an amide I band with maximum absorbance at 1621 cm–1, which is typical of a β-pleated sheet structure maintained by strong hydrogen bonds. The presence of intermolecular interactions was confirmed by the residual amide II band after 2 hours of hydrogen/deuterium exchange. Within the sequence 109–122, the hydrophobic segment AGAAAAGA (i.e., residues 113–120) was found to be central to β-sheet association. On the other hand, the α-helix promoting solvent hexafluoroisopropanol induced PrP109–122 to adopt α-helical conformation. The extension of this sequence toward the N-terminus to include the lysine residues 106 and 104 (i.e., peptides PrP106–122, 105–122, and 104–122) resulted in an increase in α-helix/random coil structure in aqueous solutions at physiological pH (Nguyen et al., 1995a). Solid-state NMR confirmed that peptide PrP109–122 can exist in at least two different conformations, depending on the solvent from which the final preparation is obtained (Heller et al., 1996). Although peptide lyophilized from acetonitrile/water solution showed β-sheet structure in the segment spanning residues 112–121, the sample lyophilized from hexafluoroisopropanol had a metastable α-helical conformation in the segment comprising residues 113–117. The expo-

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sure of lyophilized PrP109–122 to water vapor caused a complete conversion of the secondary structure to β sheet within a few hours. Evidence indicates that the physicochemical environment primarily affects the structure of the N-terminal segment of PrP109–122. X-ray diffraction of samples lyophilized from water showed that the β sheets were composed of small side chain, suggesting a single β strand in the C-terminal segment. Conversely, specimens dehydrated after acetonitrile solubilization exhibited two types of β sheets, one with larger and one with smaller side chains; this suggested the presence of two β strands corresponding to residues 109–113 and 116–122, respectively, with a turn creating an intrachain intersheet interaction (Inouye and Kirschner, 1998). It was advanced that the ionization state of His111 may play an important role in these structural modifications, as it faces the hydrophobic segment 116–122. When this residue becomes neutral by being deprotonated, the structure of the peptide may be stabilized; however, when His111 is positively charged at acidic pH, the side chain is unlikely to remain in such a nonpolar environment. Physicochemical studies of the synthetic peptide spanning residues 106–126 of human PrP led to similar conclusions. PrP106–126 consists of an N-terminal polar head (KTNMKHM-) followed by a long hydrophobic tail (–AGAAAAGAVVGGLG); and its structural features are markedly influenced by solvent composition, ionic strength, and pH. CD spectroscopy showed that the peptide adopts a random coil conformation in deionized water, a combination of random coil and β sheet in phosphate buffer pH 7.0, a predominantly β-sheet structure in phosphate buffer pH 5.0, and an α-helical structure in trifluoroethanol or in the presence of micelles formed by a 5% SDS solution. Notably, the β-sheet conformation is extremely stable, because it is not affected by trifluoroethanol if the peptide was previously suspended in phosphate buffer pH 5.0 (De Gioia et al., 1994). Studies with mutated analogs of PrP106–126 support the hypothesis that the conformational plasticity of the peptide is largely due to His111, which is located between the hydrophilic and hydrophobic segments (Salmona et al., 1999). Replacement of L-His111 with D-His111 abolished the pH-dependent conformational polymorphism and caused loss of β-sheet structure. Conceivably, the orientation of the imidazo moiety of D-His produces a steric hindrance with the neighboring Lys110 and Met112, resulting in the formation of a random coil structure. Substitution of His111 with the hydrophobic, nonionizable amino acid alanine, remarkably decreased the solubility of the peptide, which instantly formed amorphous aggregates in aqueous solutions, suggesting that hydrophobic forces prevailed over the kinetics of fibril assem-

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bly. Unexpectedly, substitution of His111 with lysine, which is always ionized in the pH range 5–7, did not abolish the pH dependent conformational polymorphism and fibrillogenic ability of the peptide. This finding is likely related to the intramolecular environment of residue 111, which is flanked by hydrophilic and hydrophobic regions that influence the pKa of lysine (Salmona et al., 1999). Determination of solution structure by NMR spectroscopy showed that PrP106–126 adopts an α-helical conformation in the hydrophobic C-terminal segment (residues 112–125) when dissolved in deionized water at pH 3.5, the most populated helical region corresponding to Ala115-Ala118. In water/trifluoroethanol, the α helix increased in population, particularly in the Gly119-Val122 tract (Fig. 4A). Conversely, in dimethyl sulfoxide, the region corresponding to the hydrophobic cluster Ala113-Ala120 adopted a prevalently extended conformation (Fig. 4B). Under all experimental conditions, the N-terminal segment spanning residues Asn108-Met112 showed a “turn-like” conformation. The analysis confirmed that the structure of PrP106–126 is remarkably sensitive to pH, supporting the view that the pH-dependent ionizable side chain of His111 may be important in modulating the conformational mobility of the peptide (Ragg et al., 1999). B. The Globular Domain Evidence suggests that the proximal part of the gobular domain extending from residue 125 to the origin of the first α helix is involved in the conformational transition of PrPC into disease-specific species. This region, which includes a short β strand, corresponds to the C-terminal segment of the amyloid protein purified from GSS brains (Fig. 3B). A synthetic peptide homologous to residues 127–147 of human PrP was found to assemble into fibrils having ultrastructural features, tinctorial properties, and x-ray diffraction patterns of amyloid (Tagliavini et al., 1993). However, the intrinsic potential of this peptide to amyloid formation appeared to be significantly lower than that of PrP106–126 and required the peptide to be intact, as shorter peptides spanning residues 127–135 or 135–147 were nonfibrillogenic. Further, the fibrils showed a distinct morphology and the propensity to helically wind around each other with a well-defined periodicity, suggesting that the sequence comprising residues 127–147 may contribute to the morphology of scrapie-associated fibrils extracted from brains of human and animals with prion diseases (Merz et al., 1981; Merz et al., 1983). Gasset et al. (1992) examined the physical characteristics of a peptide spanning residues 129–141 of hamster PrP by FTIR. The spectrum of

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FIG. 4. Solution conformations of PrP106–126 as determined by NMR and restrained molecular dynamics. Representative structures of the peptide in trifluroethanol/water solution (A) and dimethyl sulfoxide (B).

the amide I region of hydrated peptide films was marked by maximum absorbance at 1655 cm–1, consistent with multiple-ordered structures with little indication of any intermolecular interaction. Nguyen et al. (1995a) found that PrP129–141 is soluble and has coil or α-helical structures in solution; however, this peptide formed β-sheet-rich, insoluble aggregates on addition of a peptide corresponding to residues 109–122 of Syrian hamster PrP. This remarkable effect appeared to be specific, as it was not observed using unrelated amyloid peptides (e.g., Aβ11–25 and Aβ25–35) instead of PrP109–122; furthermore, the conversion process was significantly less efficient using a peptide homologous to the mouse PrP sequence, which differs from Syrian hamster PrP at residues 109 and 122 (Nguyen et al., 1995a). These findings suggested that the high propensity for β structure and fibril formation enables PrP109–122 to convert peptides having an intrinsically low amylidogenic potential from a soluble coiled state to an insoluble βsheet-rich form. Overall these data pointed to the region comprising the distal part of the flexible N-terminal domain and the proximal part of the globular domain as a possible nidus at which the conformational change is initiated in PrPC to PrPSc conversion. On this ground, larger peptides corresponding to residues 109–141 and 90–145 of Syrian hamster PrP were analyzed (Zhang et al., 1995). The latter is of particular interest because

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residue 90 is the N terminus of the protease-resistant core of PrPSc, and residue 145 coincides with a stop codon in the PRNP gene, which is associated with a cerebrovascular form of PrP amyloidosis (Kitamoto et al., 1993; Ghetti et al., 1996b). CD and FTIR spectroscopy showed that PrP90–145 adopts α-helical structure in aqueous solutions containing organic solvents such as trifluoroethanol and hexafluoroisopropanol, or detergents such as sodium dodecyl sulfate and dodecyl phosphocholine. Sodium chloride at physiological concentration or acetonitrile induced the formation of β sheets, whose intermolecular nature was evident from the presence of rod–shaped polymers as revealed by electron microscopy (Zhang et al., 1995). X-ray diffraction analysis confirmed the three-dimensional organization of these polymers, which contained β-sheet structures with intermolecular β sheets. The hydrogen-bonding distances within the β sheets were similar to those observed for hydrated prion rods, suggesting that the amyloidogenic core of PrPSc is closely modeled by the peptide PrP90–145 (Nguyen et al., 1995b). Recently, Kaneko et al. (2000) addressed the question as to whether a similar synthetic peptide is able to initiate prion disease. In particular, they used a peptide homologous to mouse PrP residues 89–143 (i.e., human residues 90–144) with the P101L substitution corresponding to GSSlinked P102L mutation. Intracerebral inoculation of a β-sheet-rich form of this peptide into transgenic mice expressing low levels of the P101L mutation resulted in neurological dysfunction and neuropathological changes consistent with GSS. Conversely, larger doses of a non β form of the same peptide failed to induce these changes, emphasizing the importance of protein conformation in disease initiation and propagation. NMR spectroscopy of recombinant murine PrP showed that helix 1 corresponding to residues 144–154 is located in the outer shell of the globular domain. Six of the 11 amino acid residues are able to change their ionization state in dependence of pH variations (Riek et al., 1996). The N-terminal half of this helix is part of the epitope of the PrPSc-specific monoclonal antibody 15B3 and is the epitope of the antibody 6H4, which recognizes both recombinant PrP and PrPC (Korth et al., 1997). On this ground, it was advanced that helix 1 might be involved in conformational changes leading to PrPSc formation. To test this hypothesis, Liu et al. (1999) examined a peptide corresponding to residues 143–158 of mouse PrP and found that it has an extraordinary high helix propensity. Comparison of NMR structure of this peptide and recombinant mouse PrP revealed striking similarities, as both formed a pincette-like motif in the N-terminal part of the helix.

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Gasset et al. (1992) investigated the secondary structure of peptides spanning residues 178–191 and 202–218 of Syrian hamster PrP (Fig. 3C). These peptides were predicted to correspond to helical regions, and indeed are part of helix 2 and helix 3, respectively, as determined by NMR (Donne et al., 1997). Unexpectedly, FTIR spectroscopy showed that hydrated PrP178–191 contained a mixture of β sheet and turns, whereas PrP202–218 had a predominantly β-sheet structure. IV. UNRAVELING THE PATHOGENESIS OF PRION DISEASES USING SYNTHETIC PEPTIDES A. Interaction of PrP Peptides with Cell Membranes A variety of data argue that plasma membranes are central to the pathogenesis of prion disease. Myoclonus, periodic electroencephalographic (EEG) changes, and spongiform degeneration of neurons, which are typical features of CJD, have been regarded as a consequence of nerve cell damage following plasma membrane abnormalities (Bass et al., 1974; Chou et al., 1980; Traub and Pedley, 1981). This view was consistent with the observation that rats injected intracortically with a membrane-bound ATPase inhibitor developed periodic EEG waves and cortical spongiosis (Bignami and Palladini, 1966). The membrane hypothesis was further supported by the demonstration that scrapie infectivity is associated with the brain membrane fraction (Millson et al., 1971; Semancik et al., 1976; Marsh et al., 1984; McKinley et al., 1991). Subsequent studies showed that cultured neurons exposed to PrPSc exhibit alteration of membrane protein, increased membrane microviscosity, and abnormal receptor-mediated calcium ion responses in the absence of morphological changes, suggesting that PrPSc affects the plasma membrane early (Kristensson et al., 1993). Caveolae-like domains purified from scrapie-infected cells and scrapie-infected brains contain both PrPC and PrPSc (Vey et al., 1996). Notably, the conversion of PrPC into PrPSc is inhibited by lovastatin, a drug that reduces membrane cholesterol levels, a critical determinant of the architecture of caveolae-like domains (Taraboulos et al., 1995, Murata et al., 1995). Accordingly, this membrane compartment has been regarded as a putative site of PrPC>PrPSc conversion. Harris and co-workers reported that mutant PrP generated by transfected CHO cells and PrPSc produced by scrapie-infected neuroblastoma cells remain associated with the plasma membrane after digestion with a specific bacterial phospholipase that

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cleaves the glycosylphosphatidylinositol (GPI) anchor. However, the anchor becomes susceptible to the enzyme when membrane proteins are denatured with SDS, suggesting that the GPI of mutant PrP and PrPSc is physically shielded from the phospholipase, either by aggregation of the protein or by intrinsic conformational features of the polypeptide chain (Lehmann and Harris, 1995; Narwa and Harris, 1999). Studies with cell-free translation systems containing ER-derived microsomal membranes have revealed that PrP may exist in different topological forms, including two transmembrane forms with opposite membrane orientation. A specific transmembrane form was found to be increased in transgenic mice expressing the A117V mutation and was detected in brain tissue of patients with GSS A117V (Hedge et al., 1998). The membrane-spanning domain of the transmembrane forms of PrP corresponds to residues 113–135. Noteworthy, synthetic peptides relevant to this domain interact with artificial and natural membranes in vitro, causing profound changes. The incubation of peptide PrP106–126 with liposomes resulted in a remarkable increase in microviscosity, as determined by steady-state fluorescence spectroscopy using the lipophylic probe 1,6-diphenyl-1,3,5-hexatriene (DPH), which incorporates into the lipid bilayer. This effect was also observed using a PrP106–126 analog with amidated C terminus (PrP106–126 NH2), a modification that substantially decreases peptide fibrillogenic ability. Conversely, a scrambled sequence of PrP106–126 and peptide PrP127–147 was not effective (Salmona et al., 1997). Similar changes in membrane microviscosity following PrP106–126 treatment were observed in several cell lines as well as in primary nerve and glial cell cultures. These changes occurred in the first few minutes of exposure and remained stable for at least 60 minutes. The increase in microviscosity was unrelated to membrane protein and lipid composition or phase transition of membrane lipids in the range of biologically significant temperatures (Salmona et al., 1997). To investigate whether this effect was due to insertion of PrP106–126 in the membrane bilayer, the peptide and its scrambled congener were coupled to a fluorescent dye and incubated for 60 minutes with murine fibroblasts. Fluorescence microscopy showed that PrP106–126 labeling was punctuated and confined to the cell surface, whereas scrambled PrP106–126 was homogeneously distributed over the cell body (Fig. 5A). When the incubation was prolonged for 24 hours, PrP106–126 was largely internalized from the cell surface to cytoplasmic organelles (Fig. 5B). These observations suggest that PrP106–126 has a high propensity to embed itself into the lipid bilayer, increasing the lipid density and

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membrane microviscosity. This propensity seems to be related to the primary structure rather than the aggregation properties of the peptide, because the poorly fibrillogenic PrP106–126 NH2, but not the fibrillogenic PrP127–147, induced similar changes. It is conceivable that the membrane modifications induced by PrP106–126 are at the basis of receptor and channel dysfunction of nerve and glial cells, and account for cell responses to the peptide in vitro. The membrane effects of PrP 106–126 were also investigated by Lin et al. (1997) using a model of planar lipid bilayer membranes. The study showed that ion-permeable channels are rapidly formed on the addition of peptide at concentrations comparable to those required for neurotoxicity. These channels were permeable to all ions and irreversibly associated with the membranes, disrupting the selectivity of cell permeability. The effects were more pronounced using an “aged” PrP106–126 and were enhanced by acidic pH, suggesting that the aggregation state of the peptide influenced channel formation. Based on the observation that a C-terminal fragment of Aβ comprising residues 29–42 is highly fusogenic (Pillot et al., 1996) and that this fragment has sequence homology with residues 118–135 of PrP, Pillot et al. (1997) investigated the interaction of peptides PrP118–135 and PrP120–133 with artificial membranes. The conformation of these peptides at the lipid/water interface was calculated by the energy minimization approach (Brasseur, 1991). PrP118–135 and PrP120–113 inserted into the lipid bilayer in an oblique way, although with different angles (25° and 45°, respectively). PrP118–135 penetrated the lipid phase through its more hydrophobic N-terminal segment, which corresponds to the hydrophobic domain of PrP106–126. Both peptides resulted highly fusogenic when exposed to unilamellar lipid vesicles and caused the leakage of entrapped calcein as well (Pillot et al., 1997). The high propensity of these peptides to insert stably into cell membranes, resulting in membrane perturbation, has recently been propsed as a mechanism of neurotoxicity for PrP peptides, which seems to be independent from the expression of endogenous PrPC (Haïk et al., 2000). B. Effects of PrP Peptides on Neuronal Cultures 1. PrP 106–126 The accumulation of PrPSc and PrP amyloid in the central nervous system is accompanied by activation of microglial cells, hypertrophy, and proliferation of astrocytes and degeneration of neurons leading to

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FIG. 5. Fluorescence pattern of murine fibroblasts after short or prolonged incubation with PrP106–126 coupled to a fluorescent dye. After one hour incubation, the fluorescent peptide is confined to the cell surface (A) while after 24 hours is internalized, the fluorescence being associated with cytoplamic organelles (B).

variable degrees of atrophy of the target regions. The temporal and topographical relationships between PrP deposition and brain changes (DeArmond et al., 1988; Bruce et al., 1989; Williams et al., 1994) suggest that altered forms of the protein may be responsible for nerve cell degeneration and glial cell reaction. To test this hypothesis, several studies have been carried out to assess the effects of synthetic PrP peptides on neurons and glial cells in culture. The first peptides to be used corresponded to consecutive segments of the GSS amyloid protein (residues 81–88, 89–106, 106–126, and 127–147). Exposure of primary rat hippocampal neurons to these peptide for 24 hours did not affect cell survival. Conversely, there was great neuronal loss in cultures after 7-day treatment with micromolar concentrations of PrP106–126. Under

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the same conditions, the other PrP peptides and a scrambled sequence of PrP106–126 did not significantly reduce cell viability (Fig. 6A and B). The neurotoxicity of PrP106–126 was dose-dependent; the toxic response was first detected at a concentration of 10 µM, was statistically significant at 25 µM, and resulted in virtually complete neuronal loss at 50 µM. Fluorescence microscopy following culture treatment with DNA-binding fluorochromes (e.g., Hoechst 33258), as well as electron microscopy revealed that neurons exposed to PrP106–126 exhibited apoptotic changes such as condensation of the chromatin and fragmentation of the nucleus. Accordingly, agarose gel electrophoresis of DNA extracted from cultured cells after 7-day treatment with the peptide showed an apoptotic pattern of DNA fragmentation, resulting from cleavage of nuclear DNA in internucleosomal regions (Forloni et al., 1993). A subsequent study revealed that the neurotoxic effect of the peptide involves calcium entry through L-type voltage-sensitive calcium channels (Florio et al., 1998). Exposure of GH3 cells—a clone derived from a GH-secreting rat pituitary adenoma—to PrP106–126 resulted in apoptosis and dose-dependent inactivation of L-type calcium channels, as determined by elctrophysiological measurements. The apoptotic pathway induced by PrP106–126 is not associated with increased expression of classic oncogenes such as C-fos, C-jun, and C-myc but is accompanied by a reduction of Bcl-2 expression (Forloni et al., 1996). Recently, we have evaluated the role of caspases in peptide neurotoxicity by measuring the activity of caspase-3 (CPP32), a major apoptosis effector enzyme, in primary cortical neurons. Culture treatment with 2.5 to 50 µM PrP106–126 resulted in dose-dependent activation of CPP32, which was 6-fold increased at the maximal dosage. Co-treatment of cultures with a nonselective caspase inhibitor (z-VAD-fmk) or a CPP32-specific inhibitor (DEVD-CHO) abolished CPP32 activation (Fig. 7A). Although z-VAD-fmk also reduced PrP106–126 neurotoxicity, co-treatment of cultures with DEVD-CHO did not prevent neuronal death, suggesting that the neurotoxic effect of the peptide was not dependent on caspase-3 activation (Fig. 7B). This observation is consistent with the immunohistochemical finding of a large disproportion between activated caspase-3 immunoreactive cells and apoptotic cells in the cerebellum of GSS patients (Migheli et al., 2000). Brown et al. (1994) reported that PrP106–126 neurotoxicity is dependent on the expression of endogenous PrPC. This finding is consistent with in vivo observations that PrP0/0 mice are resistant to scrapie infection (Büeler et al., 1993), and the presence of PrPC is essential for the development of PrPSc-related neurodegenerative changes (Brandner et al., 1996). The relationship between peptide toxicity and PrPC expres-

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FIG. 6. Effects of PrP106–126 on neurons, astrocytes, and microglia. Photomicrographs of cultures of primary rat hippocampal neurons (A and B), primary rat astroglial cells (C and D), and a human microglia cell line (E and F) after 7-day exposure to micromolar concentrations of PrP106–126 (B, D, F) or a scrambled sequence of the peptide (A, C, E). PrP106–126 treatment results in severe neuronal loss (B), hypertrophy and proliferation of astrocytes as revealed by immunohistochemical staining of glial fibrillary acidic protein (D), and activation of microglia as deduced by the change from ramified to ameboid morphology (F).

sion was further analyzed by Hope et al. (1996) who found that cell lines with low expression levels of PrPC are less susceptible to PrP106–126 than cells with high PrPC expression. Using cerebellar cultures obtained from different lines of PrP transgenic mice, Brown (1998) observed that granule cells from Tg35 mice having 8 to 10 times PrPC overex-

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FIG. 7. Relationship between caspase activation and neurotoxicity of PrP106–126. (A) The activity of caspase-3 (CPP32) was determined after 5-day exposure of cortical neurons to 50 µM PrP106–126 or scrambled PrP106–126 (Control). The remarkable increase in CPP32 activity induced by PrP106–126 was abolished by cotreatment of cultures with a specific (DEVD-CHO) or a nonselective (z-VAD-fmk) caspase inhibitor. (B) Viability of cortical neurons under the conditions described in (A). The neurotoxic effect of PrP106–126 was partially reduced by cotreatment of cultures with z-VAD-fmk, whereas DEVD-CHO was ineffective. Data are the mean ± SE of 6 determinations; *P < 0.01 versus control group (Tukey’s test).

pression were more susceptible to PrP106–126 neurotoxicity that those derived from normal mice. Unexpectedly, no significant differences were detected between granule cells from Tg20 transgenic mice having 12 to 14 times PrPC overexpression and neurons from normal mice.

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This discrepancy was explained by a different behavior of Tg35 and Tg20 microglial cells present in culture. The role of microglia in PrP106–126 neurotoxicity was first investigated by Brown et al. (1996). These authors showed that the toxic effect of the peptide to mouse cerebellar granule neurons was related to the presence of microglial cells, which responded to PrP106–126 by increasing their oxygen radical production. The neurotoxicity was remarkably reduced by treatment of cultures with L-leucine methylester (LLME), which is toxic to microglia, and was restored by addition of microglial cells. Similar results were observed using phaeochromocytoma PC12 cells (Brown et al., 1997). Treatment of undifferentiated or NGF-differentiated PC12 with 80 µM PrP106–126 did not reduce cell viability; however, when PC12 were co-cultured with microglia, the peptide induced cell degeneration, that was more pronounced after differentiation with NGF. Based on the finding that selective expression of PrPC in astrocytes is sufficient to restore susceptibility of PrP0/0 mice to scrapie infection, it was advanced that astroglial cells feature in prion disease pathogenesis, possibly by an indirect toxic effect on neurons (Raeber et al., 1997). This hypothesis is supported by an in vitro study where PrP0/0 neurons became susceptible to PrP106–126 neurotoxicity when co-cultured with PrP+/+ astrocytes (Brown, 1999). Our studies do not support the view that PrP106–126 neurotoxicity requires microglia or astrocytes. On the contrary, the presence of astrocytes in neuronal cultures may favor the neuroprotective effect of antioxidants. This was deduced by the observation that although primary neurons cultured with or without 10% fetal calf serum (FCS) are equally sensitive to the toxic effect of the peptide, only neurons cultured with FCS, a condition that markedly increases astrocyte contamination, are protected by antioxidants even in the absence of microglia (Angeretti et al., in press). The view that PrP106–126 has a direct effect on neurons is supported by the finding of peptide-induced toxicity in glia-free, neuronal cell lines such as PC12 and NB41A3 (Hope et al., 1996). The neurotoxicity of PrP106–126 has been used as a tool to identify neuroprotective compounds. Flupirtine and sulfated glycosaminoglycans have been found to prevent the toxic effects of the peptide on primary rat cortical neurons and human neuroblastoma cell lines, respectively (Perovic et al., 1995; Perez et al., 1998). Although flupirtine has been regarded as an antiapoptotic agent (Perovic et al., 1998), the effect of sulfated glycosaminoglycans was associated with the ability of these molecules to inhibit peptide aggregation and amyloid formation. However, evidence suggests that the neurotoxicity of PrP106–126 is not strictly related to the fibrillogenic properties of the peptide. Based on

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conformational studies on Aβ 25–35 (Terzi et al., 1994), we synthesized a PrP106–126 analog with amidated C-terminus (PrP106–126 NH2). Although this modification strongly reduced the ability of the peptide to form amyloid fibrils, it did not affect its neurotoxicity (Salmona et al., 1999). On the other hand, PrP106–126 NH2 lost the capability to induce astroglial proliferation, suggesting that this effect is related to aggregation state of the peptide (Rizzardini et al., 1997, Salmona et al., 1999). Jobling et al. (1999) investigated the role of the hydrophobic core sequence AGAAAGA in PrP106–126 neurotoxicity. The substitution of two or more hydrophobic residues with hydrophilic serine residues abolished the toxic effect of the peptide on mouse cerebellar neuronal cultures. This correlated with a decrease in β-sheet structure that altered the aggregation and fibrillogenic properties of the peptide. The discrepancy between this observation and our results with PrP106–126 NH2 could be due to substantial differences in the nature of the changes introduced in the peptides (i.e., alteration of the primary structure and disruption of the hydrophobic core versus removal of the C-terminus electric charge by amidation without sequence modification). The view that fibrillogenesis is not essential for neurotoxicity is supported by studies with synthetic peptides homologous to Alzheimer’s disease Aβ protein (Pike et al., 1991; Forloni et al., 1997; Johnson and Gibbs, 1998; Kshet et al., 1999). A recently published article did not confirm the neurotoxic effect of PrP106–126 and indicated that the possible source of the neurotoxic activity could be an high-performance liquid chromatography (HPLC) contaminant (Kunz et al., 1999). In this regard, it is noteworthy that the neurotoxicity of PrP106–126 has been successfully replicated in many laboratories throughout the world. Furthermore, PrP 106–126 was obtained from several different sources, both industrial and academic, and the possibility that a chemical contaminant with similar properties was systematically and exclusively associated with PrP106–126 is unrealistic (Forloni et al., 2000). 2. PrP Peptides Carrying Mutations Associated with Human Prion Diseases We have analyzed the influence of point mutations linked to CJD (D178N, E200K, I210V) and GSS (P102L, P105L, A117V, F198S, Q217R) on physicochemical properties and biological activity of PrP peptides (Foloni et al., 1999). For the study, wild-type and mutated PrP fragments corresponding to residues 89–106 (P102L and P105L variants), 106–126 (A117V), 169–185 (D178N), 195–213 (F198S, E200K and I210V variants), and 201–220 (Q217R) were synthesized. Nerve and glial cell viability was determined after prolonged exposure of rat primary neuronal and astroglial cultures to micromolar concentrations of

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peptides. Although mutations P102L and D178N strongly increased the neurotoxic activity of the native sequence, the other substitutions had no effect. The neurotoxicity of PrP89–106 carrying the P102L mutation was not related to changes in aggregation properties, because the peptide was not able to form amyloid fibrils, similar to the wild-type sequence. Conversely, the D187N substitution resulted in striking increase in fibrillogenic capacity. Furthermore, PrP169–185 with D187N mutation was able to induce astroglial proliferation, at variance with the wild-type sequence. These data suggest that PrP sequences in the mutated form may be neurotoxic and further emphasize that neurotoxicity may be independent of amyloidogenesis. C. Effects of PrP Peptides on Astroglial and Microglial Cultures Based on the observation that glial cell activation occurs early in the course of experimental prion disease and is related to PrP deposits (Williams et al., 1994), several studies focused on the effects of synthetic PrP peptides on astrocytes and microglia in vitro. Prolonged exposure of primary astroglial cultures to peptide PrP106–126 resulted in a remarkable increase in size and density of astroglial processes (Forloni et al., 1994, Fig. 4C and D). The hypertrophy of astrocytes was associated with a striking increase in glial fibrillary acidic protein (GFAP) transcripts as revealed by Northern blot analysis. Densitometric quantification of GFAP mRNA showed that this increment was dependent on peptide concentration, being significant at 10 µM and resulting in 3- and 5-fold increase above control values at 25 µM and 50 µM, respectively. The rise of GFAP transcripts was accompanied by a substantial increase in GFAP, as determined by Western blot analysis (Forloni et al., 1994). Furthermore, astrocytes exposed to PrP106–126 showed increased expression of apolipoprotein J (ApoJ). This effect was maximal after 7 days and was concentration-related, being significant at 25 µM and resulting in 3-fold increment at 50 µM (Chiesa et al., 1996). It is noteworthy that a striking increase in ApoJ mRNA has been found in experimental scrapie and ApoJ immunoreactivity is associated with PrPSc and PrP amyloid in CJD and GSS brains. Furthermore, PrP106–126 was able to enhance proliferation of astrocytes. This effect was remarkable when cultures were kept in serum-free medium. Under these conditions, the uptake of thymidine after 9-day treatment with 50 µM peptide was 50 times higher than the basal values (Florio et al., 1996) The proliferative effect of PrP106–126 was abolished by cotreatment of astroglial cultures with nicardipine, (i.e., a blocker of L-type voltage-sensitive calcium channels). Microfluorimetric analysis of intracellular calcium levels in single astrocytes showed that

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PrP106–126 induced a rapid increase in cytosolic calcium concentrations, followed by a slow return to basal levels. This effect was not observed with the scrambled peptide; it was absent when calcium was removed from the medium and was prevented by preincubation of cultures with nicardipine. These data suggest that PrP106–126 stimulates astroglial proliferation via an increase in intracellular calcium concentration, through the activation of L-type voltage-sensitive calcium channels. Brown (1999) has proposed that PrP106–126 neurotoxicity may be partly mediated by the ability of the peptide to inhibit the glutamate uptake by astrocytes. Further, it has been advanced that this inhibition may occur through the interaction of the peptide with PrPC of astrocytes based on the finding that a lower rate of glutamate uptake was related to PrPC expression (Brown and Mohn, 1999). The involvement of glutamate-mediated neurotoxicity in prion disease pathogenesis has been hypothesized by other authors (Müller et al., 1993; Scallet and Ye, 1997); however, the supposed role of PrPC in glutamate uptake needs to be further investigated. The prolonged exposure of primary mouse microglial cultures to micromolar concentrations of PrP106–126 resulted in enhanced cell proliferation and increased oxygen radical production (Brown et al., 1996). These effects were not dependent on the expression of endogenous PrPC. Treatment of a pure culture of human resting microglia with PrP106–126 induced morphological changes with a shift from ramified to ameboid morphology, enhanced phagocitic activity, and cell proliferation (Fig. 6E and F). These effects were associated with calcium entry through voltage-sensitive calcium channels and were blocked by specific inhibitors of these channels (Silei et al., 1999). Studies with neuronmicroglia co-cultures and mixed glia cultures indicate that the activation of microglia increases nerve cell death and astrocyte proliferation after peptide treatment. Nevertheless, the observations that PrP106–126 is toxic to microglia-free neuronal cell lines and induces hypertrophy and proliferation of highly purified astrocytes (Florio et al., 1996; Fleminger and Curtis, 1997) point to a direct effect of the peptide on nerve and astroglial cells, in addition to a microglia-mediated effect. Peyrin et al. (1999) found that microglial activation by PrP106–126 was associated with 5-fold overexpression of proinflammatory cytokine IL-1β and IL-6, but TNFα and anti-inflammatory cytokines IL-10 and TGF(β1) were unchanged. These effects were dependent on the ability of the peptide to form fibrils, because PrP106–126 NH2 was ineffective. These findings are in agreement with microglial activation and cytokines production observed in experimental scrapie (Williams et al., 1994; Kim et al., 1999; Baker et al., 1999) and suggests that proinflammatory cytokines may contribute to prion disease pathogenesis.

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V. CONCLUDING REMARKS The hypothesis that PrPC misfolding leads to a neurodegenerative process that can be transmitted by abnormal PrPSc conformers urged at the recognition of polypeptide segments that are central to PrPC-PrPSc interaction and PrPC>PrPSc conversion. Molecular modeling and NMR studies have suggested putative critical regions. Noteworthy, some PrP fragments from these regions were found to possess both an unusual structural polymorphism and the ability to reproduce pathological hallmarks of the disease process in vitro. Accordingly, PrP peptides can be regarded as a prime tool for investigating the molecular basis of protein misfolding and disease pathogenesis. The need to develop therapies for prion diseases has remarkably increased following the emergence of the new variant CJD. Although several compounds have been found to antagonize prion propagation in vitro or in vivo, the most suitable target for pharmacological therapy has not yet been identified. One possible strategy is to recognize compounds able to interfere with PrP domains involved in the interaction of PrPC with PrPSc or other macromolecules that feature in the conversion process. An alternative approach is to develop compounds capable of binding PrPSc selectively and of destabilizing the misfolded structure of the protein. Evidence suggests that short synthetic peptides homologous to PrP domains implicated in abnormal folding and bearing amino acid residues that destabilize β-conformation may possess these properties (Chabry et al., 1998; Soto et al., 2000). Furthermore, synthetic PrP amyloid and large PrP peptides that can be folded into different conformers can be conveniently used for large-scale screening of compounds in vitro. ACKNOWLEDGMENTS This study was supported by the Italian Ministry of Health, Department of Social Services (RF99–38), the European Community (BMH4 CT98-6011, CT98-6051) and Telethon Italia (E574). We are grateful to Dr. Thomas Stockner for the critical comments given in the preparation of the manuscript.

REFERENCES Angeretti, N., Rodrigues Martin, T., Peressini, E., Lucca, E., Galbeten, J. L., Bugiani, O., Tagliavini, F., Salmona, M., and Forloni, G. (2001). Neuroscience, in press. Baker, C.A., Lu, Z.Y., Zaitsev, I., and Manuelidis, L. (1999). J. Virol. 73, 5089–5097. Bass, N.H., Hess, H.H., and Pope, A. (1974). Arch. Neurol. 31, 174–182.

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BIOSYNTHESIS AND CELLULAR PROCESSING OF THE PRION PROTEIN BY DAVID A. HARRIS Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, Missouri 63110

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Cell Biology of PrPC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Expression and Function of PrPC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Structure and Biosynthesis of PrPC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Posttranslational Cleavage of PrPC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Subcellular Localization and Trafficking of PrPC . . . . . . . . . . . . . . . . . . . . . E. Clathrin-Coated Pits, Caveolae, and Rafts . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Cell Biology of PrPSc. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Cell Culture Models of Prion Formation. . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Subcellular Localization of PrPSc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Membrane Attachment of PrPSc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Kinetics of PrPSc Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Subcellular Site of PrPSc Formation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. edPosttranslational Cleavage of PrPSc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. INTRODUCTION Prion diseases, also called spongiform encephalopathies, are fatal neurodegenerative disorders that have attracted enormous attention not only for their unique biological features, but also for their impact on public health. This group of diseases includes kuru, CreutzfeldtJakob disease (CJD), Gerstmann-Sträussler syndrome (GSS), and fatal familial insomnia (FFI) in human beings, as well as scrapie in sheep and goats, bovine spongiform encephalopathy in cattle, and encephalopathies in mink, cats, mule deer, elk, and several exotic ungulates. Three different manifestations of prion diseases are recognized: infectious, familial, and sporadic. All three are due to the conformational conversion of a normal cellular glycoprotein (PrPC) into a pathogenic isoform (PrPSc), which is the primary component of infectious prions (Harris, 1999; Prusiner, 1999). This unusual molecular transformation is thought to involve an increase in the β-sheet content of the molecule, primarily in its N-terminal half. In infectious cases, this change is catalyzed by a sequence-specific, physical interaction between PrPC and PrPSc, in familial cases it is induced by mutations in the PrP molecule, 203 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

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and in sporadic cases it may occur spontaneously as a result of stochastic fluctuations in the structure of the protein. Defining the mechanisms underlying the generation of PrPSc from PrPC has become one of the central issues in prion research. To address this issue, it is necessary to understand something about the biosynthesis and cellular processing of both forms of PrP. This chapter reviews this subject, focusing first on PrPC and then on PrPSc.

II. CELL BIOLOGY OF PrPC A. Expression and Function of PrPC PrPC is normal cellular protein that is expressed in neurons and glia of the brain and spinal cord, and at lower levels in several peripheral tissues (Caughey et al., 1988a; Bendheim et al., 1992; Manson et al., 1992; Harris et al., 1993b; Moser et al., 1995; Dodelet and Cashman, 1998). PrP expression begins early in embryogenesis, and increases as development proceeds (Manson et al., 1992; Harris et al., 1993b). In the adult central nervous system, PrP and its mRNA are widely distributed, with particular concentrations in neocortical and hippocampal neurons, cerebellar Purkinje cells, and spinal motor neurons (Kretzschmar et al., 1986; DeArmond et al., 1987). The normal function of PrPC remains unknown, although its localization on the cell surface would be consistent with roles in cell adhesion and recognition, ligand uptake, or transmembrane signaling. Defining the physiological role of PrPC may be relevant to understanding the disease state, because the protein may fail to perform its normal function when it is converted to the PrPSc isoform. In principle, the phenotype of mice in which the PrP gene has been ablated might provide clues to the normal function of PrPC. There has been considerable confusion on this subject, as some lines of PrP-null mice display no gross developmental or anatomic defects (Büeler et al., 1992; Manson et al., 1994), whereas others exhibit a neurological illness characterized by ataxia and degeneration of cerebellar Purkinje cells (Sakaguchi et al., 1996; Moore et al., 1999). It now appears that these discrepancies can be explained by the presence of a gene for a PrP-related protein called doppel that is located downstream of the gene encoding PrP itself (Moore et al., 1999; Weissmann and Aguzzi, 1999). In the two lines of PrP-null mice with neurological abnormalities, disruption of the PrP gene causes synthesis of PrP-doppel chimeric transcripts, thereby increasing expression of doppel, which is presumably pathogenic. Although the discovery of the doppel gene provides an explanation for

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the ataxia and Purkinje cell loss seen in some lines of PrP-null mice, there is still disagreement about whether other lines with normal doppel levels and no gross neurological deficits exhibit subtle electrophysiological and structural abnormalities in the hippocampus (Collinge et al., 1994; Manson et al., 1995; Whittington et al., 1995; Colling et al., 1996; Colling et al., 1997), alterations in circadian rhythm and sleep pattern (Tobler et al., 1996), and changes in learning and memory (Nishida et al., 1997). Taken together, the studies of PrP-null mice fail to provide substantial insight into the normal function of PrPC. Several key observations have recently suggested a possible role for PrPC in metabolism of the essential trace metal, copper. First, it has been found that purified recombinant PrP, as well as synthetic PrP peptides, bind copper ions with low micromolar affinity via a series of four octapeptide repeats that reside in the N-terminal half of the protein (Hornshaw et al., 1995; Brown et al., 1997a; Stöckel et al., 1998; Viles et al., 1999). Binding is specific for copper over other transition metals, and is pH dependent, with affinity falling sharply below pH 6 (Stöckel et al., 1998; Miura et al., 1999). Of interest, copper binding causes conformational changes in the octapeptide repeats, raising the possibility that the metal may trigger some functional alteration in PrPC (Hornshaw et al., 1995; Miura et al., 1996). A second major observation is the report by one group that there is a 10- to 15-fold reduction in the content of copper, but not of several other metals, in brain membranes from PrP-null mice compared with wild-type control animals (Brown et al., 1997a). This striking result suggests that PrPC could be a major copper-binding protein in brain. Finally, the same group has reported that the enzymatic activity and copper loading of Cu-Zn superoxide dismutase (SOD1) are 50% of normal in brain and cultured cerebellar neurons from PrP-null mice, and that SOD1 activity and copper loading are elevated in mice that overexpress PrP (Brown et al., 1997b; Brown and Besinger, 1998). These observations suggest that PrP may play some role in delivery of copper to cuproenzymes such as SOD1. Our recent results (Waggoner et al., 2000) contrast with those of Kretzschmar and colleagues (Brown et al., 1997a; Brown et al., 1997b; Brown and Besinger, 1998), and require a reevaluation of the role of PrPC in copper metabolism. We find that that the amount of ionic copper in subcellular fractions from brain, measured by mass spectrometry, is similar in three lines of mice with 0, 1, and 10 times the normal level of PrPC. We also find that the enzymatic activities of Cu-Zn superoxide dismutase and cytochrome c oxidase in brain extracts are similar in these groups of animals, as is the incorporation of 64Cu into Cu-Zn superoxide dismutase both in cultured cerebellar neurons and in vivo. Our results indicate that

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PrPC is unlikely to be a major copper-binding protein in brain membranes, and also that it is probably not the primary carrier responsible for entry of copper into the brain via the blood-brain or blood-cerebrospinal fluid barriers, or for uptake of the metal into neurons from the extracellular space. Our studies also suggest that PrPC does not play a role in the specialized trafficking pathways involved in delivery of copper to SOD1 and COX in brain. A much more likely function for PrPC is to serve as a reversible sink or carrier for copper ions. Consistent with this suggestion, the normal concentration of Cu+2 in plasma and CSF (1–10 µM) is similar to the estimated Kd for copper binding to PrPC, and the concentration of the metal in brain tissue is estimated to be even higher (100 µM) (Smith, 1983). Further work will be necessary to explore whether PrPC functions as an endocytic receptor for cellular uptake of copper ions, as we have speculated (Pauly and Harris, 1998) (see later), or whether it facilitates some other aspect of copper trafficking such as efflux from the cell or intracellular sequestration of the metal. B. Structure and Biosynthesis of PrPC The mammalian PrP gene encodes a protein of approximately 250 amino acids that contains several distinct domains, including an N-terminal signal peptide, a series of five proline- and glycine-containing octapeptide repeats, a central hydrophobic segment that is highly conserved, and a C-terminal hydrophobic region that is a signal for addition of a glycosyl phosphatidylinositol (GPI) anchor (Fig. 1). PrPC is synthesized in the rough endoplasmic reticulum (ER) and transits the Golgi on its way to the cell surface. During its biosynthesis, PrPC is subject to cleavage of the N-terminal signal peptide, addition of N-linked oligosaccharide chains at two sites, formation of a single disulfide bond, and attachment of the GPI anchor (Fig. 1) (Stahl et al., 1987; Turk et al., 1988; Haraguchi et al., 1989). The N-linked oligosaccharide chains added initially in the ER are of the high-mannose type and are sensitive to digestion by endoglycosidase H; these are subsequently modified in the Golgi to yield complex-type chains that contain sialic acid and are resistant to endoglycosidase H (Caughey et al., 1989). The GPI anchor is added in the ER after cleavage of the C-terminal hydrophobic segment. It has a core structure common to other glycolipid-anchored proteins, but is modified by the addition of mannose, ethanolamine, and sialic acid residues (Stahl et al., 1992). Available evidence indicates that the oligosaccharide chains and GPI anchors of PrPC and PrPSc do not differ, although complete structures have been worked out only for PrPSc (Endo et al., 1989; Stahl et al., 1992; Stimson et al., 1999).

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FIG. 1. Structure and posttranslational processing of PrP. (Upper) Structure of the primary translation product of mammalian PrP. The five proline/glycine-rich repeats in mouse PrP have the sequence P(Q/H)GG(T/G/S)WGQ. (Lower) Structure of the mature protein. The GPI anchor attaches the polypeptide chain to the membrane. (See Fig. 4B for a schematic of the core anchor structure.) Arrows A and B indicate the positions of cleavage sites in PrPC, and arrow C a cleavage site in PrPSc. Site A lies within the GPI anchor, between the glycerolipid moiety and the ethanolamine residue that is attached to the C-terminal amino acid. Site B lies near position 110, and site C near position 89. (Reprinted with permission from Harris, 1999).

There is evidence that experimental alterations in N-glycosylation modify the biosynthetic transport of PrPC and alter its biochemical properties. Mutation of both consensus sites for N-glycosylation, or of the more N-terminal site alone, causes the protein to misfold and accumulate in a compartment proximal to the mid-Golgi stack (Rogers et al., 1990; Lehmann and Harris, 1997). However, correct N-glycosylation is not absolutely required for biosynthetic transport, since mutation of the C-terminal consensus site alone, or synthesis of wild-type PrPC in the presence of the glycosylation inhibitor tunicamycin, still allows a substantial number of molecules to be expressed on the cell surface (Lehmann and Harris, 1997). Glycosylation of PrPC is enhanced by removal of the disulfide bond, either by dithiothreitol reduction, or by mutation of the cysteine residues (Capellari et al., 1999). Surprisingly,

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molecules with either one or both N-glycosylation consensus sites mutated exhibit several biochemical properties of PrPSc, and the same is true to a limited extent for wild-type PrPC molecules synthesized in the presence of tunicamycin (Lehmann and Harris, 1997; Ma and Lindquist, 1999). Thus, wild-type PrPC has an intrinsic tendency to adopt some PrPSc-like features during its normal conformational maturation, but N-glycan chains protect against this change. The recently described doppel gene encodes a protein of 179 amino acids with 20% to 25% homology to mammalian PrP (Moore et al., 1999). This molecule represents an N-terminally truncated version of PrP that lacks the octapeptide repeats and central hydrophobic segment, but which retains three predicted α-helical regions in the C-terminal half. Doppel is expressed at high levels in testis, and at very low levels in brain. At this point, little is known about its cell biology. It is glycosylated when expressed in transfected cells, consistent with the presence of a predicted N-terminal signal sequence. In addition, there is a C-terminal hydrophobic segment that is likely to serve as a signal for GPI attachment. It will clearly be of great interest now to further investigate the properties of doppel expressed in cells and compare them to those of PrP. C. Posttranslational Cleavage of PrPC Our own work (Harris et al., 1993a) and that of others (Caughey et al., 1988b; Caughey et al., 1989; Tagliavini et al., 1992; Borchelt et al., 1993; Chen et al., 1995; Taraboulos et al., 1995) have shown that PrPC undergoes two posttranslational cleavages as part of its normal metabolism. One cleavage (labeled A in Fig. 1) occurs within the GPI anchor and releases the polypeptide chain into the extracellular medium. Many other GPIanchored proteins undergo a similar cleavage, which may result from the action of cell-surface phospholipases (Low, 1989). The second cleavage (labeled B in Fig. 1) is proteolytic and occurs within a segment of 16 hydrophobic amino acids that is completely conserved in all cloned PrP species. Our own data indicate that cleavage B occurs within an endocytic compartment of the cell (see next section), although Taraboulos et al. (1995) suggest that cholesterol-rich domains of the plasma membrane are involved. The physiological significance of the cleavages is uncertain. D. Subcellular Localization and Trafficking of PrPC PrPC is a cell-surface protein, the majority of which is found on the plasma membrane. Because it is attached by its GPI anchor, it can be released from the cell surface by treatment with the bacterial enzyme

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phosphatidylinositol-specific phospholipase C (PIPLC), which cleaves off the glycerolipid portion of the anchor (see Fig. 4B) (Borchelt et al., 1990; Caughey et al., 1990; Lehmann and Harris, 1995). There is evidence that PrPC in neurons is concentrated in the synaptic region. The protein is axonally transported to nerve terminals (Borchelt et al., 1994) and is enriched in presynaptic membranes obtained by subcellular fractionation (Herms et al., 1999). It has also been localized to synaptic profiles by immunoelectron microscopy (Fournier et al., 1995; Salès et al., 1998). By light microscopic immunocytochemistry, PrPC is found primarily in synaptic fields of the olfactory bulb, limbic structures, striatonigral complex, and cerebellar molecular layer, with little staining of neuronal perikarya or fiber pathways (Salès et al., 1998; Herms et al., 1999). These data would suggest a role for PrPC in synaptic function, although additional studies are clearly necessary. We have learned a great deal about the subcellular trafficking of PrPC from studies of transfected cell lines that express the protein. Our results demonstrate that PrPC does not remain on the cell surface after its delivery there, but rather constitutively cycles between the plasma membrane and an endocytic compartment (Fig. 2) (Shyng et al., 1993). This conclusion is based on a number of pieces of evidence, including the sensitivity of cleavage B to lysosomotropic amines, leupeptin, and brefeldin A; uptake of fluorescently labeled anti-PrP antibodies from the cell surface; and internalization and recycling of PrP molecules that have been labeled with membrane-impermeant iodination or biotinylation reagents. In cultured neuroblastoma cells, PrPC molecules cycle through the cell with a transit time of ~60 minutes, and during each passage 1% to 5% of the molecules are cleaved at site B. This endocytic recycling pathway is of interest for two reasons. First, it may be the route along which certain steps in the conversion of PrPC to PrPSc take place (see later). Second, the existence of a recycling pathway suggests that one physiological function of PrPC might be to serve as a receptor for uptake of an extracellular ligand, by analogy with the receptors responsible for uptake of transferrin and low-density lipoprotein. One attractive candidate for such a ligand is the copper ion. We have hypothesized (Pauly and Harris, 1998) that PrPC binds copper ions on the cell surface and then delivers them to an endocytic compartment within which the bound ions dissociate from PrPC and are transferred to other copper-carrier proteins that move the ions into the cytosol; PrPC would then return to the cell surface to begin another cycle. Consistent with this model, we have found that copper ions at physiologically relevant concentrations rapidly and reversibly stimulate endocytosis of PrPC from the cell surface (Pauly and Harris, 1998). It

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FIG. 2. Cellular trafficking and cleavage of PrP. After reaching the cell surface, PrPC is internalized into an endocytic compartment from which most of the molecules are recycled intact to the cell surface. A small percentage of the endocytosed molecules are proteolytically cleaved (site B in Fig. 1), and the N- and C-terminal cleavage products are then externalized. Some of the membrane-anchored protein is released into the extracellular medium by cleavage within the GPI anchor (site A in Fig. 1). (Reprinted with permission from Shyng et al., 1993).

has also recently been reported that the efficiency of radioactive copper uptake by cultured neurons is related to their expression level of PrPC (Brown, 1999). E. Clathrin-Coated Pits, Caveolae, and Rafts We have found that clathrin-coated pits and vesicles are the morphological structures responsible for endocytic uptake of PrPC (Shyng et al., 1994). This conclusion is based on immunogold localization of PrPC in these organelles by electron microscopy; inhibition of PrPC internalization by incubation of cells in hypertonic sucrose, which disrupts clathrin lattices; and detection of PrPC in purified preparations of coated vesicles from brain. The N-terminal half of the PrPC polypeptide chain is essential for efficient clathrin-mediated endocytosis, because

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deletions within this region diminish internalization of PrPC measured biochemically and reduce the concentration of the protein in coated pits determined morphometrically (Shyng et al., 1995a). The involvement of clathrin-coated pits in endocytosis of PrPC is surprising, as GPI-anchored proteins like PrPC lack a cytoplasmic domain that could interact directly with adapter proteins and clathrin. Indeed, it has been speculated that other GPI-anchored proteins are excluded from coated pits and are internalized via other surface invaginations called caveolae (Anderson, 1993). We find, however, that caveolae are not responsible for internalization of PrPC in neuronal cells, as these cells lack morphologically recognizable caveolae and do not express caveolin (Shyng et al., 1994). To explain the association of PrPC with coated pits, we have postulated the existence of a “PrPC receptor,” a transmembrane protein that has a coated-pit localization signal in its cytoplasmic domain, and whose extracellular domain binds the N-terminal portion of PrPC (Harris et al., 1996). Binding of copper ions may enhance the affinity of PrPC for this putative receptor, in line with the stimulatory effect of the metal on endocytosis of PrPC (Pauly and Harris, 1998). We have begun to search for a PrPC receptor by identifying cell-surface binding sites for radioiodinated bacterial fusion proteins incorporating segments of the PrP sequence (Shyng et al., 1995b). Thus far, we have detected saturable and specific binding sites on the surface of cultured cells that have an affinity of 70 to 240 nM, with ~1 million sites per cell. Binding is seen only with fusion proteins incorporating the N-terminal half of the PrP sequence, consistent with the importance of this region in endocytic targeting. Many, but not all of the detected binding sites are glycosaminoglycan (GAG) molecules, based on their biochemical properties. Whether either the GAG or non-GAG sites play a role in endocytosis of PrPC remains to be determined. We (Gorodinsky and Harris, 1995) and others (Naslavsky et al., 1996; Vey et al., 1996) have observed that when cells are extracted with Triton X-100 at 4°C, PrPC is recovered along with other GPI-anchored proteins in large, detergent-resistant complexes that contain cholesterol and sphingolipids, as well as signaling molecules such as src-family tyrosine kinases and G protein subunits. Complexes of similar composition have been prepared from a number of cells and tissues, and it has been speculated that they represent the biochemical equivalent of caveolae (Chang et al., 1994; Lisanti et al., 1994). Our results indicate that this hypothesis cannot be true, however, because caveolae are not found in the neuronal cells from which we have extracted PrP-containing complexes. Although it is possible that the complexes are simply an artifact

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of detergent extraction, it is has been hypothesized that they correspond to specialized microdomains of the plasma membrane (“rafts”) that exist in the intact cell and have specialized physiological functions (Simons and lkonen, 1997). Recent evidence suggests that different kinds of rafts may be present on the neuronal surface, and that PrPC may be enriched in some but not others (Madore et al., 1999). III. CELL BIOLOGY OF PrPSC A. Cell Culture Models of Prion Formation Although several cell types are thought to produce PrPSc following prion infection in vivo, only some neuronally derived cell lines appear to be susceptible to infection with scrapie prions in vitro, including N2a mouse neuroblastoma cells (Butler et al., 1988; Race et al., 1988), PC12 rat pheochromocytoma cells (Rubinstein et al., 1991), spontaneously immortalized hamster brain (HaB) cells (Taraboulos et al., 1990), and T-antigen immortalized hypothalamic neurons (GT1 cells) (Schätzl et al., 1997). Once infected, these cells continuously produce low levels of PrPSc, which can be recognized by its biochemical properties and by its infectivity in animal bioassays. Surprisingly, the infected cells display no obvious cytopathology, with the exception of the GT1 cells, a subpopulation of which appears to undergo apoptosis (Schätzl et al., 1997). Scrapie-infected N2a cells show alterations in bradykinin-mediated responses and in membrane fluidity, although these abnormalities do not seem to affect the growth or viability of the cells (Kristensson et al., 1993; Wong et al., 1996). Scrapie-infected cultured cells provide a model of infectious forms of prion diseases, but until recently there had been no comparable model of familial forms. In principle, it should be possible to model familial prion diseases by expression in transfected cells of PrP molecules carrying pathogenic mutations. By comparing cells expressing mutant PrPs with those that have been infected with exogenous prions, it should then be possible to directly contrast the familial and infectious manifestations of prion disease at the cellular level. It is important to keep in mind that the mechanisms underlying PrPSc formation may be different for the infectious and genetic forms. In the first case, exogenous PrPSc serves as a catalyst for conversion of endogenous PrPC to the PrPSc state; in the second case, mutant PrP is spontaneously transformed to PrPSc. During the last several years, our laboratory has developed a model of familial prion diseases by constructing stably transfected lines of Chinese hamster ovary (CHO) cells that express murine homologs of

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mutant PrPs associated with all three familial prion diseases of humans (Lehmann and Harris, 1995; Lehmann and Harris, 1996b; Lehmann and Harris, 1996a; Daude et al., 1997; Lehmann et al., 1997; Lehmann and Harris, 1997). We find that each of seven different PrPs carrying a pathogenic mutation displays biochemical properties of PrPSc. As shown in Fig. 3, these properties include detergent-insolubility, manifested by sedimentation at 265,000 g from Triton/deoxycholate lysates, and protease-resistance, evident by production of an N-terminally truncated core of 27 to 30 kDa after treatment with proteinase K. Wild-type mouse PrP and mouse PrP carrying a mutation (M128V) homologous to a nonpathogenic polymorphism of human PrP do not display these properties. We have noted differences among the mutant PrPs in their degree of detergent insolubility and other biochemical properties (Lehmann and Harris, 1996a; and unpublished observations). Interestingly, those molecules with the most prominent PrPSc properties are those carrying mutations that have been found to maximally decrease the thermodynamic stability of the molecule (Liemann and Glockshuber, 1999). In support of the relevance of the cell culture model to an in vivo system, we have recently demonstrated that a mutant PrP expressed in transgenic mice acquires PrPSc-like properties identical to those seen in CHO cells, and in addition produces clinical neurological dysfunction and neuropathology in the animals (Chiesa et al., 1998, 2000). There are now several other cultured cell systems that have been used to analyze the metabolism of mutant PrP molecules. We have recently shown that mutant mouse PrPs synthesized in PC12 and BHK cells have the same biochemical properties as those produced in CHO cells, and that differentiation of PC12 cells induced by nerve growth does not affect these properties (Chiesa and Harris, 2000; Harris and Ivanova, manuscript in preparation). Priola and Chesebro (1998) have reported that PrP molecules carrying CJD-linked octapeptide insertions are partially protease-resistant and detergent-insoluble when expressed in N2a neuroblastoma cells and 3T3 fibroblasts. Interestingly, these properties were more pronounced for molecules with longer insertions, and for proteins expressed in 3T3 cells compared to N2a cells. In 3T3 cells, PrP molecules with longer insertions were also more resistant to PIPLC release, although in N2a cells the mutant proteins were as releasable as wild-type PrP. In M17 human neuroblastoma cells, PrP molecules with a D178N mutation and either methionine or valine at codon 129 were found to be degraded before reaching the cell surface, a defect that was particularly severe for unglycosylated forms (Petersen et al., 1996). The mutant proteins were protease-sensitive and PIPLC-

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releasable. In these same cells, the GSS-linked Q217R mutation caused inhibition of surface transport and synthesis of forms lacking a GPI anchor, and also increased the protease resistance and detergent insolubility of the protein (Singh et al., 1997). Finally, PrP molecules carrying a Y145amber mutation linked to a variant form of GSS were found to retain their N-terminal signal sequence and were rapidly degraded by a proteasome-mediated pathway in M17 cells, suggesting that some portion of the mutant polypeptide is exposed to the cytoplasm (Zanusso et al., 1999). The Y145amber molecules were also found to be detergent-insoluble and protease-resistant.

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FIG. 3. PrPs carrying disease-related mutations are detergent-insoluble and proteaseresistant when expressed in cultured CHO cells. (A) CHO cells expressing wild-type (WT) and mutant mouse PrPs were labeled with [35S]methionine for 20 minutes and then chased for 3 hours. Detergent lysates of the cells were centrifuged first at 16,000 g for 5 minutes, and then at 265,000 g for 40 minutes. PrP in the supernatants and pellets from the second centrifugation was immunoprecipitated and analyzed by SDS-PAGE. PrP-specific bands were quantitated using a Phosphor-Imager, and the percentage of PrP in the pellet was calculated. Each bar represents the mean ± SD of values from three experiments. Values that are significantly different from WT PrP by t-test (P < 0.001) are indicated by an asterisk. PrPs carrying disease-related mutations sediment (are detergent-insoluble), whereas WT and M128V PrPs remain largely in the supernatant. Human homologs of the mutant PrPs analyzed here are associated with the following phenotypes: PG14 (9-octapeptide insertion), CJD-variant; P101L, GSS; M128V, normal; D177N/Met128, FFI; D177N/Val128, CJD; F197S/Val128, GSS; E199K, CJD. (B) CHO cells expressing each PrP were labeled for 3 hours with [35S]methionine, and chased for 4 hours. Proteins in cell lysates were either digested at 37°C for 10 minutes with 3.3 µg/mL of proteinase K (+ lanes), or were untreated (– lanes), before to recovery of PrP by immunoprecipitation. Five times as many cell-equivalents were loaded in the + lanes as in the – lanes. Molecular weight markers are in kilodaltons. PrPs carrying pathogenic mutations yield a protease-resistant fragment of 27 to 30 kDa, whereas WT and M128V PrPs are completely degraded. In separate experiments, we have shown that the PrP 27–30 fragments are N-terminally truncated after the octapeptide repeats, the same region within which PrPSc from infected brain is cleaved (Lehmann and Harris, 1996b). (Modified with permission from Harris and Lehmann, 1997).

Taken together, these studies of cultured cells make it clear that pathogenic mutations induce significant alterations in the metabolism of PrP. One effect of the mutation is that the protein acquires PrPSc-like biochemical properties, although the extent to which this occurs can vary with the cell type and with the mutation being expressed. It thus seems likely that these culture systems are modeling important features of the PrPC→PrPSc conversion process. However, it is important to point out that mutant PrPs synthesized in cultured cells differ in at least one respect from PrPSc isolated from the brains of humans and animals: the cell-derived PrPs have considerably lower degree of protease-resistance. It remains to be determined whether this difference reflects a variation in protease-resistance that is often seen in scrapie isolates from different sources (Bessen and Marsh, 1992), or results from the inefficiency of cultured cells in carrying out some critical step in the conversion process. Clearly, the definitive test of whether mutant PrPs synthesized in cultured cells represent authentic PrPSc will be to determine whether these proteins are infectious in animal bioassays.

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B. Subcellular Localization of PrPSc The subcellular distribution of PrPSc has been difficult to determine, primarily because this form of the protein displays poor immunoreactivity unless treated with denaturing agents that have a deleterious effect on cell morphology. Immunofluorescence studies of scrapieinfected N2a cells suggest that some PrPSc molecules reside intracellularly, colocalizing with Golgi markers in some clones, but not in others (Taraboulos et al., 1990). In electron microscopic studies of both N2a cells and brain, PrPSc also colocalizes with late endosomal and lysosomal markers (McKinley et al., 1991; Arnold et al., 1995). It is also clear that some PrPSc molecules, both mutant and infectious, are present on the cell surface, as shown by electron microscopic immunogold staining, and by labeling of intact cells with membrane-impermeant probes (Stahl et al., 1990c; Caughey and Raymond, 1991; Jeffrey et al., 1992; Lehmann and Harris, 1996a). The tentative conclusion from these studies is that PrPSc may be widely distributed within infected cells, but further studies are clearly required. C. Membrane Attachment of PrPSc The precise mechanisms by which PrPSc is attached to cell membranes has been a matter of uncertainty. Chemical analysis of the purified protein demonstrates that PrPSc, like PrPC, possesses a C-terminal GPI anchor (Stahl et al., 1992). Unlike PrPC, however, PrPSc is not releasable by PIPLC from brain membranes or from the surface of scrapie-infected N2a cells (Caughey et al., 1990; Stahl et al., 1990b; Safar et al., 1991; Lehmann and Harris, 1996a). The GPI anchor of purified and denatured PrPSc is cleavable by PIPLC (Stahl et al., 1990a), suggesting that the anchor is not modified, for example by acylation of the inositol ring, so as to make it intrinsically PIPLC-resistant. Taken together, all of these results argue that PrPSc is associated with membranes in a way that is different from PrPC, although exactly how it differs has been unclear. Our analysis of mutant PrPs in cultured CHO cells has shed new light on the issue of the membrane attachment of PrPSc (Lehmann and Harris, 1995; Narwa and Harris, 1999). We find that mutant PrPs, like PrPSc from infected brain, are not released from membranes by treatment with PIPLC (Fig. 4A). This property does not result from absence of a GPI anchor structure, because the mutant PrPs metabolically incorporate the anchor precursors [3H]ethanolamine, [3H]palmitate, and [3H]stearate. Although we originally postulated that mutant PrPs pos-

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FIG. 4. PrPs carrying disease-related mutations are not released from the cell surface by PIPLC. (A) CHO cells expressing wild-type (WT) and mutant mouse PrPs were biotinylated with the membrane-impermeant reagent sulfo-biotin-X-NHS at 4°C, and were then incubated with PIPLC at 4°C before lysis. PrP in the PIPLC incubation media and cell lysates was immunoprecipitated, separated by SDS-PAGE and visualized by developing blots of the gel with HRP-streptavidin and enhanced chemiluminescence. PrP bands from three separate experiments were quantitated by densitometry, and the amount of PrP released by PIPLC was plotted as a percentage of the total amount of PrP (medium + cell lysate). Each bar represents the mean ± SD. Values that are significantly different from WT PrP by t-test are indicated by single (P < 0.01), and double (P < 0.001) asterisks. All of the PrPs carrying pathogenic mutations are less PIPLC-releasable than WT and M128V PrPs. E199K PrP is more releasable than the other mutants, consistent with our observation that there are subtle biochemical differences among the mutant proteins. Phenotypes associated with the homologous human PrPs are given in the legend to Fig. 3. (B) Schematic of the membrane attachment of wild-type PrPC, which can be completely released from cells by treatment with PIPLC. The core structure of the GPI anchor, along with the site cleaved by PIPLC are indicated. C-E are schematics showing how PIPLC is proposed to interact with mutant PrP. (C) On intact cells, the mutant protein adopts a PrPSc-like conformation that physically blocks access of PIPLC to the GPI anchor. It is also possible that aggregation of mutant PrP molecules contributes to the inaccessibility of the anchor. (D) After extraction from the membrane using nondenaturing detergents such as Triton X-100 and deoxycholate (DOC), the abnormal conformation of the mutant protein is maintained, and the anchor is still inaccessible to PIPLC. (E) After denaturation in SDS, the conformation of the mutant protein is disrupted, and the anchor becomes susceptible to PIPLC cleavage. (Reprinted with permission from Harris, 1999).

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sessed a secondary mechanism of membrane attachment in addition to their GPI anchors (Lehmann and Harris, 1995), more recent evidence suggests that the mutant molecules are resistant to PIPLC release because their GPI anchors become physically inaccessible to the phospholipase as part of their conversion to the PrPSc state (Narwa and Harris, 1999) (Fig. 4C-E). This conclusion is based on failure of PIPLC to quantitatively remove [3H]palmitate label from the proteins, or to render them hydrophilic by Triton X-114 phase partitioning. Resistance to cleavage is observed when PIPLC is applied to intact cells (Fig. 4C), as well as when treatment is carried out after lysis in nondenaturing buffers (Fig. 4D). However, denaturation in SDS renders the GPI anchor of the mutant PrPs susceptible to cleavage, suggesting that PIPLC-resistance depends on the native structure of the protein (Fig. 4E). We now view PIPLC-resistance as being an operational property analogous to protease resistance and postulate that it reflects an alteration in the structure of PrP attendant on conversion to the PrPSc state. Although most PrPC molecules are attached to the cell membrane exclusively by their GPI anchor, it has been reported that there exists a subpopulation that displays a transmembrane orientation. This conclusion was originally based on analysis of PrP molecules synthesized in vitro from synthetic mRNA using either wheat germ extract or rabbit reticulocyte lysate in the presence of canine pancreatic microsomes (Hay et al., 1987; Lopez et al., 1990). To explain the existence of transmembrane forms, Lingappa and colleagues proposed that a 24 amino acid region of PrP (“stop transfer effector”) induces a pause in translocation of the polypeptide chain, and that cytosolic factors determine whether this intermediate is either converted into a stable transmembrane species or is fully translocated into the ER lumen (Yost et al., 1990; De Fea et al., 1994). In subsequent work, these authors postulated the existence of two transmembrane species of PrP, each with the same membrane-spanning segment (residues 112–135), but with opposite orientations of the polypeptide chain (Hegde et al., 1998a; Hegde et al., 1998b). It has recently been suggested that one of these species (CtmPrP, which has its C-terminus in the ER lumen) is the primary effector of neurodegeneration in both inherited and infectious cases of prion diseases, as mice carrying certain mutations in the transmembrane region, as well those infected with scrapie, showed increased amounts of CtmPrP (Hegde et al., 1999). In contrast, our results suggest that CtmPrP is unlikely to be part of a general pathogenic pathway, because diseaseassociated mutations outside the transmembrane domain do not increase the amount of this form (Stewart and Harris, 2001).

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D. Kinetics of PrPSc Production The kinetics of PrPSc production have been examined both in scrapieinfected N2a and HaB cells, and in CHO cells expressing mutant PrPs. Generation of protease-resistant PrP in infected cells occurs only during the chase period after pulse-labeling, with half-maximal accumulation requiring several hours (Borchelt et al., 1990; Caughey and Raymond, 1991; Borchelt et al., 1992). This result is consistent with the idea that transformation of PrPC into PrPSc is a posttranslational event. Once formed, PrPSc appears to be metabolically stable for as long as 24 to 48 hours, in contrast to PrPC, which turns over with a half-life of 4 to 6 hours (Borchelt et al., 1990; Caughey and Raymond, 1991; Borchelt et al., 1992). Only a minority of PrPC molecules are converted to PrPSc in infected cells, with the remainder being degraded by pathways similar to those found in uninfected cells; these pathways have not been identified, but they may include acidic compartments such as endosomes and lysosomes (Taraboulos et al., 1995). We have used transfected CHO cells to identify intermediate biochemical steps in the conversion of mutant PrPs to the PrPSc state (Daude et al., 1997). Our strategy was to measure the kinetics with which three PrPSc-related properties (PIPLC resistance, detergent insolubility, and protease resistance) develop in pulse-chase labeling experiments. This has allowed us to define three steps in the conversion process (Fig. 5). The earliest biochemical change we could detect in mutant PrP, one that was observable within minutes of pulse-labeling cells, was the acquisition of PIPLC resistance, a property that was revealed by partitioning of the phospholipase-treated protein into the detergent phase of Triton X-114 lysates, or by its binding to phenylSepharose. The second step is acquisition of detergent insolubility, which is not maximal until 1 hour of chase (Lehmann and Harris, 1996b), arguing that it occurs after the acquisition of PIPLC resistance. Detergent insolubility presumably reflects aggregation of PrP molecules, and by sucrose gradient fractionation we were able to detect aggregates ranging in size from 4S (monomeric) to more than 20S (> 30 PrP molecules). The third step is acquisition of protease resistance, which is not maximal until several hours after labeling (Lehmann and Harris, 1996b). We have hypothesized that the fundamental conformational change that underlies conversion of mutant PrPC into PrPSc is reflected in the acquisition of PIPLC resistance, with detergent insolubility and protease resistance being secondary properties that develop some time after the initial molecular conversion.

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FIG. 5. A scheme for transformation of mutant PrPs to a PrPSc state. Mutant PrPs are initially synthesized in the PrPC state and acquire PrPSc properties in a stepwise fashion as they traverse different cellular compartments. PIPLC resistance, which develops in the ER, reflects folding of the polypeptide chain into the PrPSc conformation. Detergent insolubility and protease resistance, which develop on arrival at the plasma membrane or along an endocytic pathway, result from intermolecular aggregation (“maturation”). The times given underneath the boxes indicate when after pulse-labeling the corresponding property is detected. Addition of brefeldin A (BFA) to cells or incubation at 18°C, treatments that block movement of proteins beyond the Golgi apparatus, inhibit acquisition of detergent insolubility and protease resistance, but not PIPLC resistance. (Adapted with permission from Daude et al., 1997).

Although a PIPLC-resistant intermediate underlies conversion of mutant PrPs to the PrPSc state, such an intermediate may not play a role in PrPSc formation from wild-type PrPC after infection with exogenous prions. Consistent with this idea, there is evidence that PrPSc in infected neuroblastoma cells is synthesized from a precursor that is PIPLC-sensitive (Caughey and Raymond, 1991). However, much further work remains to characterize the molecular intermediates in PrPSc formation in infected cells. E. Subcellular Site of PrPSc Formation Surprisingly little information is available about how extracellular PrPSc is taken up by cells during the initial stage of infection. If this were to occur via an endocytic mechanism, then PrPSc may interact with PrPC on the plasma membrane or in endosomes, and key events in the conversion process may take place in these locations. In fact, several pieces of evidence suggest that an endocytic pathway is involved in generation of PrPSc in scrapie-infected N2a and HaB cells, including the localization of at least some PrPSc molecules in endosomes and lysosomes (McKinley et al., 1991), the incorporation of radiolabel into PrPSc after iodination of PrPC on the cell surface (Caughey and Raymond, 1991),

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and inhibition of PrPSc production either by removal of surface PrPC using PIPLC or proteases (Caughey and Raymond, 1991; Borchelt et al., 1992) or by blockage of surface delivery using brefeldin A (Taraboulos et al., 1992). These results do not distinguish between the plasma membrane and endosomes as the relevant sites for PrPSc production. Recent studies suggest that detergent-resistant subdomains of the plasma membrane (“rafts”, discussed previously) may be involved in formation of PrPSc. Consistent with this idea, both PrPC and PrPSc are found in raft domains isolated biochemically (Gorodinsky and Harris, 1995; Taraboulos et al., 1995; Naslavsky et al., 1996; Vey et al., 1996; Naslavsky et al., 1999). In addition, pharmacological depletion of cellular cholesterol, which is known to disrupt rafts, inhibits PrPSc formation (Taraboulos et al., 1995), whereas sphingolipid depletion, which does not alter the raft localization of PrP, actually enhances PrPSc production (Naslavsky et al., 1999). Finally, artificially constructed transmembrane forms of PrP, which are excluded from rafts, are poor substrates for conversion to PrPSc (Kaneko et al., 1997). Our kinetic studies of mutant PrPs synthesized in CHO cells suggest that individual steps in formation of PrPSc may take place in at least two different cellular locations (Fig. 5). Because mutant PrPs become PIPLCresistant within minutes of synthesis in pulse-labeling experiments, this early step must take place in the ER. Consistent with this conclusion, acquisition of PIPLC resistance is not affected by treatment of cells with brefeldin A or by incubation at 18°C, manipulations that block exit of proteins beyond the Golgi (Daude et al., 1997). In contrast, detergent insolubility and protease resistance, which do not develop until later times of chase, and are reduced by brefeldin A and 18°C incubation, are likely to be acquired after arrival of the protein at the cell surface, either on the plasma membrane itself or in endocytic compartments. Raft domains may be involved in these changes (unpublished data). Thus, our studies of cells expressing mutant PrPs are consistent with experiments on scrapie-infected cells, but with the additional finding that the earliest step in PrPSc synthesis—formation of a PIPLC-resistant, protease-sensitive intermediate—takes place in the ER rather than on the plasma membrane or in endosomes. Further work is required to determine whether an early ER intermediate is also generated in infected cells. If so, this would presumably require that external PrPSc somehow gain access to the lumen of the ER to interact with newly synthesized PrPC. The increase in CtmPrP observed in scrapie-infected brain (Hegde et al., 1999) is consistent with interaction between PrPSc and nascent PrPC occurring in the ER. The idea that the generation of PrPSc from mutant PrPs may begin in the ER is theoretically appealing

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because of the well-known role of this organelle in protein folding. It is also reasonable to suggest that ER chaperones are good candidates for the hypothetical cellular cofactors that are widely thought to play an important regulatory role in prion synthesis (Welch and Gambetti, 1998). A recent paper suggests another possible location for generation of PrPSc—the cytoplasm. Ma and Lindquist (1999) expressed a form of mammalian PrP that lacked N- and C-terminal signal sequences in the yeast Saccharomyces cerevisiae and found that the protein became detergent-insoluble and protease-resistant. Treatment of N2a neuroblastoma cells with dithiothreitol and tunicamycin also produced a PrPSc-like species, leading the authors to suggest that perhaps the initiating event in PrPSc formation is retrotranslocation of PrP molecules from the ER into the reducing and deglycosylating environment of the cytoplasm. F. Posttranslational Cleavage of PrPSc PrPSc in cultured cells and brain undergoes a proteolytic cleavage that removes a portion of the N-terminus (Caughey et al., 1991; Taraboulos et al., 1992; Chen et al., 1995). This cleavage occurs at a site (labeled C in Fig. 1) that is distinct from the one at which PrPC is cleaved (labeled B). Cleavage at site C ensues within an hour after formation of PrPSc and occurs within the region attacked by proteinase K to yield PrP 27–30. Sensitivity to lysosomotropic amines and protease inhibitors indicates that cleavage of PrPSc occurs in endosomes or lysosomes, compartments that may play a role in the generation of this isoform, as discussed previously. However, N-terminal trimming is not an essential step in formation of PrPSc, as PrPSc is still produced in the presence of lysosomotropic amines that inhibit the trimming (Caughey et al., 1991; Taraboulos et al., 1992). In many cases of GSS, PrPSc fragments smaller than 27 to 30 kDa have been observed that are generated by additional cleavages (Tagliavini et al., 1991; Tagliavini et al., 1994; Piccardo et al., 1995; Parchi et al., 1998). A 7 to 8 kDa fragment is produced by cleavage near residues 80 and 150, and an 11 kDa fragment by cleavage near residues 50 and 150. In some cases, these smaller fragments have been shown to be major constituents of the amyloid plaques that are characteristic of GSS (Tagliavini et al., 1991; Tagliavini et al., 1994; Piccardo et al., 1995), and it is reasonable to propose that PrP molecules carrying GSS-linked mutations are metabolized in a distinctive way to generate these amyloidogenic products.

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IV. CONCLUSIONS The work reviewed here has begun to provide a detailed picture of the biosynthesis and cellular processing of PrPC and PrPSc. PrPC is synthesized and matures along the secretory pathway much like other membrane glycoproteins. One important feature is the addition of a GPI anchor that serves to attach the polypeptide chain to the lipid bilayer without the necessity for a transmembrane domain. Once it reaches the plasma membrane, PrPC is constitutively endocytosed via clathrin-coated vesicles, and a portion of the molecules is proteolytically cleaved in a highly conserved domain before recycling to the cell surface. Efficient endocytosis depends on structural features in the Nterminal half of the PrP molecule and may be mediated by a transmembrane “PrPC receptor”.

FIG. 6. Model of the cellular pathways involved in generation of PrPSc. In the infectious manifestation of prion diseases, extracellular PrPSc in the form of a prion particle (1) interacts with PrPC on the cell surface, possibly in detergent-resistant rafts, catalyzing its conversion to PrPSc (2). Conversion may also occur after uptake of the proteins into an endosomal compartment (3). Once formed, some PrPSc accumulates in lysosomes (4), although the protein is probably found in a number of other cellular locations as well. In familial prion disorders, mutant PrPC is converted spontaneously to the PrPSc state via a series of biochemical intermediates, the earliest of which is a PIPLC-resistant form generated in the ER (5). Mutant PrP molecules are subsequently delivered to the cell surface, where they become detergent-insoluble (6) and then protease-resistant (7), possibly in raft domains. Steps 6 and 7 could also occur in endocytic organelles. (Reprinted with permission from Harris, 1999).

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The cellular pathways involved in formation of PrPSc are summarized in Fig. 6. During infectious transmission of prion disease, interaction between exogenous PrPSc and endogenous PrPC may take place in detergent-resistant rafts on the plasma membrane or in endocytic organelles, although other locations are not ruled out. Once generated, PrPSc is metabolically stable and becomes localized partly but not exclusively in intracellular organelles, perhaps endosomes and lysosomes, that may be the sites where the N-terminus of the protein is proteolytically cleaved. In familial forms of prion diseases, mutations cause stepwise changes in the biochemical properties of PrP as it is converted to PrPSc. The earliest recognizable change is acquisition of PIPLC resistance, which takes place in the ER, and may register the initial conformational alteration of the protein. Detergent insolubility and protease resistance develop later, possibly in raft domains on the plasma membrane or in endocytic compartments. Several important avenues for future investigation are suggested by the cell biological results summarized here. These include defining the mechanism by which PrPC molecules are concentrated in coated pits and vesicles, further exploring the interaction between PrPC and copper ions at the cellular level, isolating individual conversion intermediates along the pathway from PrPC to PrPSc, and identifying cellular chaperone molecules that play a role in the conversion process. Information on each of these subjects will allow us to develop a more detailed understanding of the pathogenesis of prion diseases, and will ultimately provide the basis for development of rational therapeutic modalities. ACKNOWLEDGMENTS Work in the author’s laboratory is supported by grants from the National Institutes of Health, the American Health Assistance Foundation, and the Alzheimer’s Association.

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INTERACTION OF PRION PROTEINS WITH CELL SURFACE RECEPTORS, MOLECULAR CHAPERONES, AND OTHER MOLECULES BY SABINE GAUCZYNSKI, CHRISTOPH HUNDT, CHRISTOPH LEUCHT, AND STEFAN WEISS* Laboratorium für Molekulare, Biologie-Genzentrum-Institut für Biochemie der LMU München, D- 81377 Munich, Germany

I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Cell Surface Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The Role of a Cellular Prion Protein Receptor . . . . . . . . . . . . . . . . . . . . . . B. A 66 kDa Membrane Protein as a Potential Prion Receptor . . . . . . . . . . . . C. The 37 kDa Laminin Receptor Precursor (37 kDa LRP) . . . . . . . . . . . . . . D. The Cadherins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Molecular Chaperones of Mammals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Heat-Shock Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Protein X. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Chemical Chaperones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Interaction between Prion Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Other PrP Interacting Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. PrP Ligands (PLi’s) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Bcl-2. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Laminin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Therapeutics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Nucleic Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

229 231 231 235 237 242 243 245 247 248 248 251 251 253 254 255 262 262

I. INTRODUCTION The prion protein PrP represents a central player in transmissible spongiform encephalopathies (TSEs), also known as prion diseases (for review see Lasmézas and Weiss, 2000)). The physiological role of the cellular isoform of PrP termed PrPc is speculative so far (for review see (Weissmann, 1996)) and might involve control of circadian activity rhythms and sleep (Tobler et al., 1996), maintenance of cerebellar Purkinje cell (Sakaguchi et al., 1996), and normal synaptic functions (Collinge et al., 1994; Fournier et al., 1995; Kitamoto et al., 1992). Because several reports do not describe any phenotype for PrP (Bueler et al., 1992; Lledo et al., 1996; Manson et al., 1994), the only proved role of PrPc is its necessity for the development of TSEs (Bueler et al., 1993) * Stefan Weiss is the corresponding author. 229 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

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such as bovine spongiform encephalopathy (BSE) in cattle, new variant Creutzfeldt-Jakob disease (nvCJD) in humans, or scrapie in sheep. A recent report describes a superoxide dismutase (SOD) activity for PrPc (Brown et al., 1999), suggesting that PrP might play a role in the cellular resistance to oxidative stress. In the last 20 years of the twentieth century, researchers worldwide were eagerly searching for molecules able to interact specifically with the prion protein in the hope of identifying interactors (1) that play an important role in the life cycle of prions or (2) that could be developed into powerful TSE therapeutics. This chapter summarizes PrP interacting molecules that might be relevant for PrP pathogenesis or TSE therapy. In the first section we describe putative prion protein receptors, including the role of heparan sulfate proteoglycans (HSPGs). A cellular model will be presented that describes the possible role of prion receptors and prion proteins, including the recently identified PrP-like protein termed doppel (Moore et al., 1999). The model emphasizes the possible role of PrP and its receptor regarding PrP internalization as well as signal transduction and physiological function, in particular, the 37 kDa laminin receptor precursor (LRP), an up to now unidentified 66 kDa cell surface protein, and cadherins, which are then discussed as prion receptors that might trigger the entry of PrP into scrapie infectable cells. Next, we summarize the role of molecular chaperones, including chemical chaperones that may catalyze or hamper the conversion process of PrPc to PrPSc. In this context, we emphasize a possible function for protein X, an as yet unknown protein predicted by S.B. Prusiner to be necessary for the PrP conversion process. The occurrence of PrP dimers under native and denaturing conditions observed in different cell systems and in vitro represents another aspect of PrP interactions, in this case an interaction of PrP with itself. The possible role of such PrP dimers in the complex scenario of PrP oligomerization and multimerization processes is discussed. In section V we report on a series of PrP interacting molecules identified using different biochemical approaches such as ligand blotting and yeast two-hybrid techniques. Among these are the PrP ligand proteins (Pli) encompassing Pli 3–8, Pli 45, and 110 as well as Bcl-2, which belongs to a family of proapoptopic and antiapoptopic molecules. The role of Bcl-2 in the light of neurodegeneration and apoptosis is discussed. The interaction between laminin and PrP-mediating neuritogenesis is also reported. The last section describes molecules, mainly of nonproteinaceous origin, which act as therapeutics for the treatment of TSEs. These include polyanions such as heteropolyanion 23, dextran sulfate 500, pentosan polysulfate (SP54), and heparin. Other groups of anti-TSE therapeutics include Congo red, polyene antibiotics such as AmB and MS-8209, IDX, porphorins, phtalo-

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cyanes, and the protein clusterin. The possible modes of action of these molecules such as interfering with the PrPc/PrPSc conversion process followed by PrP accumulation, interfering with the cellular uptake of PrPc/PrPSc, overstabilization of PrPSc, or competing with cellular glycosaminoglycans for binding to PrPc are discussed. The last group of PrP interacting molecules represent nucleic acids including RNA aptamers, the latter as a possible tool for the diagnosis of TSEs. II. CELL SURFACE RECEPTORS A. The Role of a Cellular Prion Protein Receptor To understand the pathogenesis of diseases such as TSEs, it is necessary to clarify how the biological system works under physiological conditions. The main principle of the “protein-only” hypothesis is that the cell-membrane glycoprotein PrPc is converted into its pathogenic isoform PrPSc, a process that involves conformational changes of the protein (Prusiner et al., 1998). During this transformation PrP acquires additional regions of β sheets in the polypeptide chain, resulting in a partial resistance to proteases. The cellular pathway of PrPc is of major interest because here the conversion of PrPc to PrPSc might take place. PrPc is synthesized in the rough endoplasmatic reticulum (rER). It is passaged via the Golgi and secretory granules to the cell surface where it is anchored to the plasma membrane by its glycosyl phosphatidylinositol (GPI) moiety (Rogers et al., 1991). According to an endocytic recycling pathway, the surface-PrPc is internalized by clathrin-coated pits (Shyng et al., 1994) or caveolae-like domains (CLDs) (Vey et al., 1996). The endocytosis of PrPc could be mediated by a transmembrane protein, which might connect the GPIanchored PrP to clathrin. Harris postulated the existence of an endocytic PrP-receptor that carries a coated-pit localization signal in its cytoplasmic domain and whose extracellular domain binds the N-terminal part of PrPc (Harris, 1999; Harris et al., 1996). He observed that deletions within the N-terminal region of PrPc result in a decrease of internalization of the protein and consequently in a reduction of the PrPc concentration in coated pits (Harris, 1999; Shyng et al., 1995). In addition, Harris observed that chicken PrP binds to the surface of mammalian cells via heparan sulfates on the cell surface (Shyng et al., 1995). Several researchers described an interaction between heparan sulfates and PrP (Brimacombe et al., 1999; Caughey et al., 1994; Chen et al., 1995; Gabizon et al., 1993). Heparan sulfates have been shown to be a component of amyloid plaques in prion diseases (Gabizon et al., 1993). Recently, it has been demonstrated that the addition of heparin competes with the binding of

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copper to PrP, which occurs in the octarepeat region (Brown et al., 1997) (Brimacombe et al., 1999), suggesting that this region of PrP binds to heparin. The recently observed superoxide dismutase (SOD) activity of PrPc is dependent on the presence of the octarepeat region (Brown et al., 1999), confirming the important role of this domain for PrP. HSPGs make up proteoglycan moieties consisting of proteins carrying glycosaminoglycan (GAGs) chains made of anionic polysaccharide chains. Heparan sulfate, the main GAG-constituent of HSPGs, like heparin, consists of disaccharide repeating units of O-/N-sulforyl and N-acetylglucosamine (or N-acetylgalactosamine) and O-sulforyliduronic acid except that it harbors fewer N- and O-sulfate groups and more N-acetyl groups. The proteoglycans HSPGs are thought to play an important role on the cell surface within the life cycle of prions. The process by which exogenous PrPSc enters the cell is unclear so far. The uptake of the infectious agent could also be mediated by a receptor protein or might occur receptor independent. The conversion of PrPc to PrPSc may take place after internalization in cellular compartments such as endosomes, lysosomes, or endolysosomes. This conversion process is thought to be influenced by an unknown protein termed protein X (Telling et al., 1995), which could represent a molecular chaperone such as Hsp60 (Edenhofer et al., 1996). In addition, it has been suggested that several proteins possessing a GPI-anchor are excluded from coated pits and internalized by caveolae (Anderson, 1993). Furthermore, it has been reported that PrPc and PrPSc are present in CLDs isolated from scrapie-infected neuroblastoma cells and brains of scrapieinfected hamsters, and it is speculated that the conversion of PrPc into PrPSc could also take place in these compartments (Vey et al., 1996). To understand the mechanism of this conversion event as well as the physiological function of the cellular prion protein, it is important to investigate the involvement of a possible receptor protein as well as of proteins showing biological properties similar to PrP, such as the recently discovered PrP-like protein designated doppel (Dpl) (Moore et al., 1999). The discovery of doppel does not only represent the first PrP-related protein (Moore et al., 1999), it also could explain some curious, surprising observations within several lines of Prnpo/o mice, which differ only in the strategy used to generate PrPc-deficiency. Creating an internal insertion or deletion within the PrP exon 3, two lines of mice were generated showing normal development without any pathological phenotype (Bueler et al., 1992; Lledo et al., 1996; Manson et al., 1994). However, in two other cell lines the entire coding sequence of PrP as well as a ~1 kb region 5′ to exon 3 including the exon 3 splice acceptor site were deleted (Sakaguchi et al., 1996). These Prnpo/o mice showed progressive symptoms of ataxia and Purkinje cell degeneration in the cerebellum. It is sug-

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gested that Dpl is involved in a physiological process in a manner leading to this pathological phenotype. Doppel is the first PrP-like protein to be described in mammals (Moore et al., 1999). It consists of 179 amino acid residues showing ~25% identity with all known prion proteins. The Dpl locus, Prnd, is located 16 kb downstream of the PrP gene, Prnp, generating two major transcripts of 1.7 and 2.7 kb. Like PrP, Dpl mRNA is expressed during the embryogenesis but, in contrast to PrP, it is poorly expressed in the adult central nervous system (CNS) and at high levels in the testis of mice. However, Dpl is upregulated in the CNS of the two Prnpo/o lines that develop late-onset ataxia and Purkinje cell death but not in the normally developed Prnpo/o lines (Moore et al., 1999). Therefore, it was assumed that Dpl may provoke neurodegeneration in PrPdeficient mice, an observation that might explain why some lines of Prnpo/o mice develop cerebellar dysfunction and Purkinje cell death, whereas others do not. Moore et al. suggested that Dpl and PrP may share some biological functions owing to the similarities between these two proteins (Moore et al., 1999). Would it be possible that PrP and Dpl bind to each other or would it also be possible that they compete for binding to a common receptor? Dpl synthesis is thought to occur in the secretory pathway to yield a globular, N-glycosylated, membrane-associated protein comparable to PrPc, but in contrast to it containing no octarepeat region in its N-terminal domain (Moore et al., 1999). In addition, expression of moderate levels of N-terminal truncated PrP with deletions of amino acid residues 32–121 or 32–134 caused ataxia and specific degeneration of the granular layer of the cerebellum in PrPo/o mice, whereas mice expressing shorter truncations of PrP, up to residue 106, show no pathological changes (Shmerling et al., 1998). This granule cell dysfunction was completely abrogated by introducing a single copy of a wild-type murine PrP gene into mice. It is speculated that the truncated PrP may compete with some other molecule with a function similar to that of PrP for a common ligand or receptor. It was assumed that in wild-type mice PrP interacts with a presumed receptor promoting signal transduction (Fig. 1A), and the same signal is elicited by interaction of the receptor with π, a conjectural protein that has the functional properties of PrP, but is not closely related to it on the DNA level (Fig. 1B) (Shmerling et al., 1998). This would explain why the absence of PrPc has no obvious phenotypic consequences. It is postulated that truncated PrP can interact with the receptor without giving rise to a signal (Fig. 1C). The affinity of the receptor for truncated PrP would have to be stronger compared to π, but would be less compared to intact PrP. Only N-terminal truncated PrP where the deletion extends to or beyond residue 121 shows cerebellar dysfunction leading to the conclusion that the globular domain of cellular PrP binds to a receptor, whereas

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FIG. 1. Model of PrPc- and receptor-mediated signal transduction. In the normal cell, PrPc and receptor molecules from the same cell or from different cells can interact and promote signal transduction (A). The same signal might be elicited by the binding of a conjectural protein designated π, which possesses the functional properties of PrPc explaining why some lines of PrPO/O mice develop normally (B). In the absence of PrPC, N-terminal truncated PrP can also interact with the receptor competing with the binding of π, however, without giving rise to a signal and leading to ataxia and degeneration of the granular layer of the cerebellum. A similar event is thought to take place in PrP-deficient mice, which are showing a pathological phenotype. In these mice a PrPlike protein called doppel (Dpl) is upregulated in the CNS. It is speculated that this protein may bind with higher affinity to the receptor than π does, resulting in ataxia and degeneration of Purkinje cells (C).

the flexible tail of the N-terminus spanning residues 23 to 120 is responsible for activation (Shmerling et al., 1998). One possible interpretation for the pathological phenotype caused by the expression of N-terminal truncated PrP is that such PrP-mutants assumes a Dpl-like conformation that is neurotoxic and results in the killing of the granular layer in the cerebellum (Moore et al., 1999). The association of Dpl overexpression with degeneration of Purkinje cells, which were rescued by overexpression of wild-type PrP, suggest that Dpl and PrP interact perhaps directly or indirectly by competing as ligands for a common receptor. Therefore, both proteins may play a role in cell contact processes (Fig. 1). Recently, a signal transduction activity of the prion protein by achieving tyrosine kinase Fyn was described (Mouillet-Richard et al., 2000). Since PrPC locates GPI-anchored at the cell surface, whereas Fyn-kinase is associated with the inner plasma membrane of the cell, a transmembrane receptor might mediate the PrPC dependent activation of the Fyn-kinase.

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In this section we describe the different candidates, identified so far, that may act as prion protein receptors. Distinct strategies and methods were used to identify the putative receptor molecule. Further investigations are necessary to clarify the identity of a physiological PrPC-receptor and to reveal its role in the normal cellular process of PrPC, as well as in the pathogenesis of prion diseases. Identification and characterization of this receptor are also important in designing drugs that could be used to prevent the initial uptake of the infectious agent into cells. B. A 66 kDa Membrane Protein as a Potential Prion Receptor Employing complementary hydropathy, a 66 kDa membrane protein that could act as a cellular prion protein receptor, was recently identified (Table I) (Martins et al., 1997). By means of this strategy, a hypothetical peptide mimicking the receptor binding site should bind to the neurotoxic domain of prion proteins. Here a peptide encoded by the DNA strand complementary to that of the human PrP gene, spanning amino acid residues 114 to 129, was chemically synthesized and used to immunize mice to generate antibodies directed against this complementary prion peptide. The available mouse antisera were used to investigate the localization of the putative receptor by immunofluorescence and confocal microscopy approaches, resulting in the detection of an antigen at the cell membrane of primary mouse neurons. In Western blot analysis of membrane extracts from mouse brain, the antiserum recognized a specific protein of 66 kDa. In vitro and in vivo binding assays were performed demonstrating that PrPC and the 66 kDa membrane protein could bind to each other (Martins et al., 1997). Flow cytometry studies revealed that purified membrane extracts, prepared from mouse brain, inhibited in vivo recognition of cellular PrP in cultured neuroblastoma cells (N2a) by anti-PrP antiserum. This process could be reversed by pretreatment of such membrane extracts with antiserum raised against the complementary prion peptide and the putative receptor protein. Furthermore, both the complementary prion peptide and the antiserum against it were able to block the neurotoxic effects mediated by the human prion peptide 106–126 toward cultured neuronal cells. Martins et al. suggested that a specific receptor for prion proteins could be responsible for their internalization and for the cellular responses mediated by PrPC. They speculated that, as PrPC tends to accumulate in postsynaptic vesicles (Askanas et al., 1993), both PrPC and its receptor are involved in interneuronal cell adhesion causing neuronal networking (Martins et al., 1997). According to Martins et al., in the normal cell PrPc and receptors from the same cell or from different cells can interact and mediate signal transduction, triggering their physiological function. They postu-

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TABLE I PrP Binding Proteins, Identity, and Characteristics PrP Binding Protein

cDNA Identified

Pli45c Pli1 10c Pli3c Pli4c Pli5c

Yes Yes Yes Yes Yes

Pli6c

Yes

Pli7c

Yes

Pli8c 37-kDa laminin receptor precursora 66-kDa proteina Cadherinsa

Yes Yes

Known Homology

Surface Protein

Method of Identification

Reference

No No No No No

ligand blot ligand blot PrP-AP screening PrP-AP screening PrP-AP screening

Oesch et al., 1990 Oesch et al., 1990 Yehiely et al., 1997 Yehiely et al., 1997 Yehiely et al., 1997

Yes

PrP-AP screening

Yehiely et al., 1997

No

PrP-AP screening

Yehiely et al., 1997

No Yes

PrP-AP screening yeast-two-hybrid screening

Yehiely et al., 1997 Rieger et al., 1997

Yes

complementary hydropathy PrP-AP screening

Martins et al., 1997

No

GFAP PSF human ESTs None guinea pig organ of corti, rat and human ESTs mouse Aplp 1 (amyloid precurser like protein) mouse Nrf2 (p45 NF-E2 related factor) none 37 kDa lamininreceptor precursor none

Yes

Cadherins

Yes

Bcl2c

Yes

Bcl-2

No

Chaperonsb

Yes

several molecular chaperons

No

yeast-two-hybrid screening various methods

Cashman and Dodelet, 1997 Kurschner and Morgan, 1995 DebBurmans et al., 1997 Edenhofer et al., 1996 Tatzelt et al., 1996

See Section II. See Section III. cSee Section V. a b

lated that the infectious agent should interact with the same receptor following internalization, facilitating the conversion of PrPc into PrPSc and leading to PrPSc accumulation and finally cell death (Martins, 1999). Further investigations leading to the identification of the 66 kDa protein are necessary to clarify the role of this putative receptor in the normal process of PrPc, as well as in the pathogenesis of TSEs.

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C. The 37 kDa Laminin Receptor Precursor (37 kDa LRP) In a yeast two-hybrid screen, we identified a specific molecule as an interaction partner for the prion protein: the 37 kDa laminin receptor precursor (37 kDa LRP) (Table I) (Rieger et al., 1997). We speculated that this protein could act as a potential receptor for the cellular PrP. This interaction was confirmed by coinfection and cotransfection studies in insect and mammalian cells, respectively (Rieger et al., 1997). Furthermore, investigations of the LRP level in several organ and tissues of scrapie-infected mice and hamsters demonstrated that LRP occurs in higher amounts only in those organs that exhibit infectivity and PrPSc accumulation such as brain, spleen, and pancreas compared with uninfected control animals (Rieger et al., 1997). This was confirmed by cell culture experiments demonstrating an increased amount of LRP in scrapie-infected mouse neuroblastoma (N2a) cells compared with uninfected cells. Mapping of the 37 kDa LRP with different peptide fragments identified a transmembrane domain containing amino acids 86–101 (Castronovo et al., 1991b) and a laminin-binding domain comprising amino acids 161–180 (Castronovo et al., 1991b), which is thought to be directed toward the extracellular space (Fig. 2). Mapping of the LRP/PrP interaction site performed in the yeast two-hybrid system demonstrated that the laminin-binding domain can also function as a PrP binding site (Rieger et al., 1997) (Fig. 2). LRP is thought to be the precursor of the 67 kDa laminin receptor (67 kDa LR) because attempts to isolate the gene for the 67 kDa LR resulted in identification of a cDNA fragment that encoded a 37 kDa polypeptide (Rao et al., 1989; Yow et al., 1988). This was confirmed by pulse-chase experiments carried out with antibodies directed against the 37 kDa protein (Castronovo et al., 1991a; Rao et al., 1989). The 67 kDa laminin receptor was first isolated from tumor cells (Lesot et al., 1983; Malinoff and Wicha, 1983; Rao et al., 1983) owing to its high binding capacity to laminin, a glycoprotein of the extracellular matrix that mediates cell attachment, movement, differentiation, and growth (Beck et al., 1990). Engelbreth-Holm-Swarm (EHS) laminin (Beck et al., 1990), which has been proved to bind to the 37 kDa LRP (Rieger et al., 1997) (Table II), consists of three polypeptide chains: A or α (440 kDa), B1 or β, and B2 or γ (each 220 kDa), linked via disulfide bonds, resulting in the typical cross-structure (Beck et al., 1990). Several other classes of laminin-binding proteins have been described including integrins (Albelda and Buck, 1990) and β-galactoside binding lectins such as galectin-3 (Bao and Hughes, 1995; Ochieng et al., 1993; Yang et al., 1996) equivalent to CBP-35 (Laing et al., 1989). Immunoblotting assays

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FIG. 2. Schematic view of the prion protein (PrP) and the 37 kDa laminin receptor precursor (LRP) on the surface of a scrapie-infectable cell. PrP is anchored by GPI (Blochberger et al., 1997) and is thought to colocalize with LRP. The putative transmembrane region of LRP stretches from aa 86 to aa101 (Castronovo et al., 1991b). The laminin binding domains from aa 161 to 180 (Castronovo et al., 1991b) encompassing the palindromic sequence LMWWML, which appeared during evolution from the nonlaminin-binding ribosomal protein p40 (Ardini et al., 1998), to the laminin-binding LRP on the cell surface is identical to the PrP binding domain (Rieger et al., 1997).

performed with a polyclonal serum directed against galectin-3 revealed that the 67 kDa LR carries galectin-3 epitopes, whereas the 37 kDa LRP does not (Buto et al., 1998). The 37 kDa LRP/67 kDa LR is a multifunctional protein (Table II), and its amino acid sequence is well conserved throughout evolution, showing a high degree of homology among mammalian species (Rao et al., 1989). The evolutionary analysis of the sequence identified as the laminin-binding site [which we proved to correspond to the PrP binding domain (Rieger et al., 1997)] suggested that the aquisition of the laminin binding capability is linked to the palindromic sequence LMWWML, which appeared during evolution concomitantly with laminin binding (Ardini et al., 1998). This protein evolved from the ribosomal protein p40, which participated in protein synthesis on 40 S ribosomes without any laminin-binding activity (Auth and Brawerman, 1992) to a cell surface receptor binding laminin (Rieger et al., 1997), elastin (Hinek et al., 1988; Salas et al., 1992), and carbohydrates (for

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TABLE II Characteristics of the 37 kDa Laminin Receptor Precursora (LRP)/67 kDa Laminin Receptorb (LR) Characteristics Isolation

Occurrence of the 37 LRP/p40 gene

Cellular localization of 37 kDa LRP

Molecular weight Binding partners of –37 kDa LRP

–67 kDa LR

Functional domains

Functions of –37 kDa LRP

–67 kDa LR

37 kDa LRP/p40 cDNA (Rao et al., 1989; Yow et al., 1988); 67 kDa LR isolated from solid tumors (Lesot et al., 1983; Malinoff and Wicha, 1983; Rao et al., 1983) Sacchoromyces cerevisiae (Davis et al., 1992), Arabidopsis thaliana (Garcia-Hernandez et al., 1994), Drosophila melanogaster (Melnick et al., 1993), Urechis caupo (Rosenthal and Wordeman, 1995), Chlorohydra viridissima (Keppel and Schaller, 1991), Haloarcula marismortui (Ouzonis et al., 1995), Candida albicans (Lopez-Ribot et al., 1994), mammals (Ardini et al., 1998) At the cell surface of mosquito cells (Ludwig et al., 1996), of Candida albicans (Lopez-Ribot, 1994), and of mammalian cells such as Madin-Darby canine kidney cells (MDCK) (Salas et al., 1992); in the cytoplasm on 40S ribosomes (Auth and Brawerman, 1992); in the nucleus (Sato et al., 1996) 37,000 (laminin receptor precursor protein) 67,000 (mature laminin receptor protein) Laminin (Rieger et al., 1997), PrPc (Rieger et al., 1997), the Venezuelan equine encephalitis (VEE) virus (Ludwig et al., 1996); association of LBP c/p40 with histones H2A, H2B and H4 (Kinoshita et al., 1998) Laminin (Beck et al., 1990), elastin and carbohydrates (for review: (Ardini et al., 1998; Mecham, 1991; Rieger et al., 1999)), the Sindbis virus (Wang et al., 1992) Transmembrane domain: aa 86–101 (Castronovo et al., 1991b), Laminin binding domain: aa 161–180 (Castronovo et al., 1991b); PrPc binding domain: aa 157 and 180 (Rieger et al., 1997) Receptor for laminin (Rieger et al., 1997), PrPc (Rieger et al., 1997) and the Venezuelan equine encephalitis virus (Ludwig et al., 1996); as ribosomal protein LRP/p40 involved in protein synthesis (Auth and Brawerman, 1992); possible role of LBPc/p40 in maintenance of nuclear structures (Kinoshita et al., 1998) Receptor for laminin (Beck et al., 1990), elastin, carbohydrates (for review: (Ardini et al., 1997; Mecham, 1991; Rieger et al., 1999) and the Sindbis virus (Wang et al., 1992); crucial role in the metastatic potential of solid tumors (Castronovo et al., 1991b)

Laminin receptor precursor, LRP. Lamin receptor, LR c Laminin binding protein, LBP (equivalent to LRP). a b

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review see (Ardini et al., 1998; Mecham, 1991; Rieger et al., 1999)). In addition, interaction of the epitope-tagged laminin-binding protein LBP/p40 with nuclear structures was observed in cultured cells (Sato et al., 1996). In vitro analysis revealed that LBP/p40 binds tightly to chromatin DNA through association with histones H2A, H2B, and H4, suggesting that this protein may play an essential role in the maintenance of nuclear structures (Kinoshita et al., 1998). The laminin receptor family is highly conserved in a wide spectrum of eukaryotic cells (Keppel and Schaller, 1991; Wewer et al., 1986), including yeast (Demianova et al., 1996), and is encoded by archaean genomes (Ouzonis et al., 1995). 37 kDa LRP acts as a receptor for the Venezuelan equine encephalitis virus on mosquito cells (Ludwig et al., 1996), whereas the 67 kDa LR functions as a receptor for the Sindbis virus on mammalian cells (Wang et al., 1992) (Table II). The mechanism of how the 37 kDa precursor protein forms the mature 67 kDa isoform is still unclear. Homodimerization of the 37 kDa LRP (Landowski et al., 1995) or the involvement of an additional component (Castronovo et al., 1991a) has been discussed. Recent studies suggested that the 67 kDa LR is a heterodimer stabilized by fatty acid-mediated interactions (Buto et al., 1998). Very recently, it has been proved that the 67 kDa LR (also termed laminin binding protein, p67 LBP) is expressed on a subset of activated human T lymphocytes and, together with the integrin, very late activation antigen-6, mediates strong cellular adherence to laminin (Canfield and Khakoo, 1999). In summary, the 37 kDa LRP/67 kDa LR polymorphism remains a mystery. Both forms may act as the receptor for prions on the surface of scrapie infectable cells. Mammalian genomes contain multiple copies of the LRP gene, in particular 6 copies in the mouse and 26 copies in the human genome (Fernandez et al., 1991; Jackers et al., 1996a), a fact that has hampered the identification of the active gene for a long time. To date, only the genes for the chicken and the human gene encoding LRP have been isolated (Clausse et al., 1996; Jackers et al., 1996b). The gene encoding 37 kDa LRP belongs to a multicopy gene family and contains seven exons and six introns (Jackers et al., 1996b). The 37 kDa LRP/p40 gene has been identified in different species, including Saccharomyces cerevisiae (Davis et al., 1992), Arabidopsis thaliana (Garcia-Hernandez et al., 1994), Drosophila melanogaster (Melnick et al., 1993), the sea urchin Urechis caupo (Rosenthal and Wordeman, 1995), Chlorohydra viridissima (Keppel and Schaller, 1991), the fungus Candida albicans (Lopez-Ribot et al., 1994) and the archaebacterium Haloarcula marismortui (Ouzonis et al., 1995), as well as in mammals (Ardini et al., 1998; for review, Rieger et al., 1999).

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The 37 kDa LRP also acts as a receptor for alphaviruses such as the Venezuelan equine encephalitis (VEE) virus on the surface of mosquito cells (Ludwig et al., 1996), has been identified on the cell surface of the fungus Candida albicans (Lopez-Ribot et al., 1994) and has been proved to be located on the surface of Madin-Darby canine kidney (MDCK) cells from dogs, which might be involved in cell attachment, spreading and polarization (Salas et al., 1992). These findings clearly demonstrate the location of the 37 kDa LRP on the cell surface. Within the life cycle of prions, LRP may play a role in the physiological function of PrPc, as well as in the pathogenesis of prion diseases. We assume that LRP is involved in the internalization process of PrPc via cavolae-like domains (Vey et al., 1996) or clathrin-coated pits (Shyng et al., 1994) (Fig. 3). Involvement of clathrin-coated pits in the endocytosis of a GPI-anchored protein such as PrPc is surprising because PrPc has no cytoplasmic domain that can interact directly with the intracellular components of coated pits (Harris, 1999). Here a receptor protein could be responsible for making the connection between the surface-anchored PrP to clathrin. The uptake of PrPSc is thought to be mediated directly by a receptor protein such as LRP, but could also be mediated in an indirect manner dependent on the presence of cellular PrP. We assume that internalized PrPSc interacts with PrPc during the endocytic pathway (Fig. 3). PrPc is probably converted to PrPSc within the endosome, lysosomes, or endolysosome influenced by an unknown protein termed protein X (Telling et al., 1995), which could represent a molecular chaperone such as Hsp60 (Edenhofer et al., 1996). Recently, a homology of the amino terminus of LRP with members of the Hsp70 family was observed (Ardini et al., 1998), suggesting that LRP/p40 might be involved in protein folding. Although we demonstrated a specific interaction between PrP and members of the Hsp60 family including GroEL (Edenhofer et al., 1996), no binding of PrP to members of the Hsp70 family was observed, which suggests no homology to the Hsp60 family (Edenhofer et al., 1996). However, it cannot be excluded that a hypothetical chaperone activity of LRP might be involved in the PrPc/PrPSc conversion reaction, which is thought to occur in endosomes, lysosomes, or endolysosomes of the endocytic pathway in the life cycle of prions. Other proteins encompassing an GPI-anchor were internalized by caveolae (Anderson, 1993). It has been suggested that PrPc and PrPSc are internalized by CLDs, a compartment where the conversion of PrPc to PrPSc might also take place (Vey et al., 1996). PrPSc accumulation leads to neuronal cell death resulting in vacuolization and death of the organism. The role of LRP within the life cycle of prions mediating PrP internalization and its involvement in pathological mechanisms within the

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FIG. 3. Model of the life cycle of prions. PrPC is synthesized in the rough endoplasmatic reticulum (ER), and after passing through the secretory pathway including the Golgi and secretory vesicles, reaches the surface of a PrPSc infectable cell where it is anchored via a glycosylphosphatidyl inositol (GPI) moiety. Endocytosis of PrPC and possibly PrPSc via clathrin coated vesicles could be mediated by the 37 kDa laminin receptor precursor (LRP). The uptake of the infectious agent could also be LRP independent. The conversion of the internalized PrPC to PrPSc is thought to take place in the endosomes, lysosomes, or endolysosomes. Molecular chaperones could be involved in this conversion process. PrP replication and aggregation can occur in neuronal cells of the brain but also in the cells constituting the lymphoreticular system. Alternatively, endocytosis and conversion of PrPC into PrPSc could happen in caveolae-like domains (CLDs).

complex scenario of transmissible spongiform encephalopathies has to be further investigated. D. The Cadherins Two cell surface proteins were isolated from murine cells and characterized as so-called prion protein binding proteins (PrPBPs) (Table I) (Cashman and Dodelet, 1997). Mouse and human PrPs expressed as fusion proteins to human placental heat-stable alkaline phosphatase (PrP-AP) bound with high affinity to the surface of many primary cells and cell lines, particularly to the mouse muscle cell line G8, whereas no binding of AP alone could be observed. Frog oocytes showing little or no intrinsic PrP-AP surface binding were microinjected with in vitro transcribed mRNA generated from pooled plasmid clones of a G8 cDNA

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library. Following selection of clones that showed specific binding to PrP-AP, sequence analysis revealed the cDNA inserts in two clones, one encoded a portion of protocadherin-43 spanning amino acid residues 67 to 252 and exhibited the highest level of PrP-AP binding activity, the other one encoded a portion of OB-cadherin-1 (the N-terminal cadherin repeat) and showed a moderate PrP-AP binding (Cashman and Dodelet, 1997). Protocadherin-43 described by Sano et al. (1993) and OB-cadherin-1 described by Okazaki et al. (1994) belong to a group of cell adhesion proteins designated Cadherins. Cadherins are a family of transmembrane glycoproteins involved in Ca2+ dependent cell-cell adhesion that occurs in many tissues mediating development patterning and tissue organization. They contain a large N-terminal extracellular region consisting of repetitive subdomains including the Ca2+-binding sites. Ca2+-binding is required for cadherin interaction and cell-cell adhesion, a process that results from lateral clustering of cadherin cis dimers and their trans association with cis dimers on the apposed cell (Steinberg and McNutt, 1999). The C-terminus consists of a transmembrane region and a highly conserved cytoplasmic domain through which cadherins interact with intracellular adhesions proteins such as catenins and stabilize the internal structure of the cell. Binding of PrP-AP to cultured cells was significantly reduced in the presence of the calcium chelator EDTA, indicating that for optimum binding, the presence of divalent cations such as Ca2+ might be required. Binding of mouse, human, and bovine cellular PrP as well as PrPSc from BSE-affected brain to the candidate receptor was observed (Cashman et al., 1999). Prion proteins could act as novel ligands for cadherin proteins. Cadherins participate in cell-layer segregation and morphogenesis in development, also in maintenance of cell-cell recognition in mature tissues, and may participate in disorders in which recognition is deficient, such as metastatic cancer. It is also possible that they are involved in muscle and immunological disorders as well as in neurodegenerative diseases such as TSEs (Cashman and Dodelet, 1997). The possible role of cadherins as cell surface receptors for prion proteins, however, has still to be confirmed. III. MOLECULAR CHAPERONES OF MAMMALS The crucial event in prion diseases involves the conformational change of the cellular form of the prion protein into the pathogenic isoform. This change causes a dramatic alteration within the structure. Structural

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variations of a protein often require a catalysing agent. Molecular chaperones are prominent candidates that could promote this reaction. The protein-only hypothesis indicates that the scrapie form of the prion protein can promote the conversion of the cellular form. This leads to the conclusion that prions themselves can act as chaperones (Liautard, 1991). Thermokinetic analysis of protein folding shows that a misfolded chaperone gives rise to new misfolded chaperones, which fit very well to the protein-only hypothesis in which PrPSc triggers the formation of PrPSc. Besides this theory, other proteins can act as promotors for the prion conversion reaction. In 1996 chemical reagents were investigated and were shown to affect formation and propagation of PrPSc. Cellular osmolytes and proteinaceous chaperones were tested in this context (Tatzelt et al., 1996b). Chaperones that can prevent the formation of PrPSc (Fig. 4) might act as powerful tools for the generation of anti-TSE therapeutics. Molecular chaperones also represent a biochemical and mechanistical link between the mammalian prions and the “prion-like” proteins in

FIG. 4. Influence of molecular and chemical chaperones on the conversion process of PrPC to PrPSc. Molecular chaperones such as Hsp 104 and GroEL promote the conversion reaction, whereas the chemical chaperones TMAO, DMSO, and sucrose prevent PrPSc formation.

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yeast. In this light heat-shock protein Hsp104 has an effect on the conversion of hamster PrP (DebBurman et al., 1997) and on the regulation of the yeast nonchromosomal element [PSI+] (Chernoff et al., 1995), suggesting that the prion concept is of general importance in mammalian and nonmammalian systems. Studies on the transmission of human prion proteins to transgenic mice indicate the existance of an unknown protein termed “protein X”, which binds to PrP (Telling et al., 1995) and might act as a molecular chaperone. A. Heat-Shock Proteins A number of cellular proteins function in vivo as chaperones that catalyse the formation of proteins with an intact secondary, tertiary, and quaternary structure. Heat shock proteins (Hsps) are prominent representatives of these chaperones and were first discovered because of their specific induction during the cellular response to heat shock (Gething and Sambrook, 1992). Nevertheless, the majority of the Hsps are expressed constitutively, and their functions are diverse. Hsps stabilize unfolded protein precursors, rearrange protein oligomers and dissolve protein aggregates in an ATP-dependent manner. Hsps are thought to play an important role in the conversion of the cellular prion protein PrPc to the pathogenic isoform PrPSc (Table III). In 1995 the expression levels of Hsp72, Hsp28, and Hsp73 in normal and scrapie-infected mouse neuroblastoma cells were investigated (Tatzelt et al., 1995). After heat shock Hsp72 and Hsp28 were both detectable in normal, but not in scrapie-infected, cells. The constitutively expressed Hsp73, however, was expressed at comparable levels in both cell types, indicating that Hsp73 could possibly assist the formation of PrPSc. The lack of Hsp72 and Hsp28 in scrapie-infected cells suggests that chaperones do not catalyse a refolding of PrPSc into PrPc in these cells. Together, both facts might lead to an increase of PrPSc concentrations in scrapie-infected cells. We identified Hsp 60 as a PrP binding molecule employing a HeLa cDNA library in prey and hamster PrP in bait position of the yeast twohybrid system (Edenhofer et al., 1996). In vitro binding studies with recombinant PrP confirmed the specificity of the PrP-Hsp60 interaction. Mapping analysis employing a series of PrP peptides identified the C-terminus of PrP (aa 180 to aa 210) encompassing α-helix 2 and parts of α-helix 3 (179–193 and 200–217)(Riek et al., 1996; Donne et al., 1997; Riek et al., 1997) as the Hsp60 binding domain on PrP. GroEL, the prokaryotic homolog of Hsp 60 revealed the same binding domain as

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TABLE III Function of Heat-Shock Proteins and Their Effect on the Prion Protein Heat Shock Protein

Reference

Hsp28

Tatzelt et al. (1995)

Hsp40

DebBurman et al. (1997)

Hsp60

Edenhofer et al. (1996)

Hsp70

DebBurman et al. (1997)

Hsp72

Tatzelt et al. (1995)

Hsp73

Tatzelt et al. (1995)

Hsp90

DebBurman et al. (1997)

Hsp104

DebBurman et al. (1997)

GroEL

Edenhofer et al. (1996) and DebBurman et al. (1997)

GroES

DebBurman et al. (1997)

First Reported Function in Prion Diseases

Effect on PrP Conversion

Role in Ca2+-dependent thermoresistance

No effect on PrP conversion/PrPSc diminishes synthesis of Hsp28 Co-chaperone of Hsp70s No effect on PrP conversion Stabilization of prefolded Binding to haPrP, structures and folding binding domain: aa 180–210 Completion of transNo influence on PrP location in mitochondria conversion Prevents aggregation and No effect on PrP accelerates refolding of conversion/PrPSc damaged proteins diminishes synthesis of Hsp72 Cytosolic heat shock Assists PrPSc formation? protein Stabilizing of inactive pre- No influence on PrP cursor forms in the conversion cytosol Thermotolerance and Promotes conversion of ethanol tolerance in PrPc yeast Antifolding before Binding to haPrP, translocation binding domain: aa 180–210, promote conversion of PrPC form functional complex No influence on PrP with GroEL conversion

Hsp60 on PrP. This indicates that eukaryotic as well as prokaryotic chaperones interact with the prion protein and suggest an important role for heat shock proteins in the conversion process of prion proteins. GroEL and the heat shock protein Hsp104 are able to affect the in vitro conversion of hamster PrP, confirming the importance of GroEL for the PrP conversion reaction (DebBurman et al., 1997). However, this process requires the presence of exogenous added PrPSc, suggesting that the conversion process and further aggregation seem to require a nucleation seed. Molecular chaperones may not be sufficient for this reaction. Other heat shock proteins such as GroES, Hsp40,

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Hsp70, and Hsp90 do not show any effect in the conversion process. Hsp104 links mammalian prion proteins and the prion-like yeast protein Sup35. Hsp104 could thereby either promote sup35* or sup35 formation depending on Hsp 104 concentrations. Hsp 104 might influence the regulating process of the [PSI+] element in S. cerevisiae (Patino et al., 1996). In conclusion, heat shock proteins might influence the structure of mammalian and yeast prions. B. Protein X The transmission of human prion proteins to transgenic mice depends on the species of the endogenous expressed transgenic prion protein and the homozygocity/heterocygocity status of the expressed transgene. In contrast to transgenic mice ablated for the mouse Prnp gene or transgenic mice expressing low levels of a chimeric transgene, which are susceptible toward human prions, transgenic mice expressing the human PrP transgene are completely resistant toward human prions. This phenomenon reflecting the species barrier can be explained by a species specific factor termed protein X, which is thought to participate in prion formation. Protein X might act as a chaperone facilitating or hampering the conversion of PrPc to PrPSc. The fact that transgenic mice hyperexpressing human PrP are resistant to human prions (Telling et al., 1995), together with the finding that transgenic mice expressing chimeric MHu2MPrPC retain human PrP susceptibility suggests that protein X could bind to the cellular form of the prion protein and the affinity of protein X to prion proteins of different species may vary. The binding of protein X to the prion protein may result in the PrP conversion reaction. Differences in the amino acid sequence of PrP of different species may be the main reason for both effects. The main differences between mouse and human PrP are thought to reside in the carboxy-terminus of PrP. An epitope mapping of the binding site for protein X on PrP (Kaneko et al., 1997b) by substitution of the basic residues at aa position 167, 171, or 218 preventing PrPSc formation suggests that the binding site for protein X on PrP resides within this region. Amino acid 218 is located within the third α helix of the mouse prion protein and residues 167 and 171 reside within an adjacent loop. The stoichiometry of the protein X/PrPc complex is unknown to date. The fact that the protein X/PrPc interaction was abolished by mutations preventing the PrPSc formation might be useful for the development of anti-TSE therapeutic agents. A prerequiste for that, however, is the identification of protein X.

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C. Chemical Chaperones In contrast to “classical” chaperones consisting of proteins, chemical chaperones represent chemical compounds of small molecular weight that are able to stabilize proteins and correct misfolded ones (Welch and Brown, 1996)(Fig. 4). Chemical chaperones such as glycerol, trimethylamine-N-oxide (TMAO), and dimethyl sulfoxide (DMSO) might stabilize the native conformation of a protein by direct interaction. These compounds termed “cellular osmolytes” are produced in cells in response to osmotic shock (Somero, 1986). Glycerol, TMAO, and DMSO were tested to determine their influence on the formation of PrPSc in ScN2a cells (Tatzelt et al., 1996b). All reduced the extent of PrP conversion into its detergent insoluble form. The stabilizing effect of the native form of a protein was also demonstrated for other proteins such as the cystic fibrosis transmembrane regulator (CFTR) (Brown et al., 1996). The presence of chemical chaperones might have an effect on the hydration of proteins. Because self-association or tighter packaging of the prion protein is enhanced, PrPSc fails to interact with PrPc so that no PrPc/PrPSc heterodimer is formed leading to an inhibition of the PrP conversion process (Gekko and Timasheff, 1981). In the case that chemical chaperones might be transported to the brain bypassing the blood-brain barrier (BBB), they might be useful as therapeutic agents in TSE-therapy. The influence of chemical chaperones has also been demonstrated in cell-free conversion assays (DebBurman et al., 1997). The conversion of hamster PrP using partially denatured PrPSc was only inhibited by DMSO. Glycerol and cyclodextrin compounds had no effect, whereas molecular chaperones (Hsp104) were able to block the conversion process. Chemical chaperones such as glycerol and cyclodextrin, acting as co-chaperones, might have an influence on molecular chaperones that are lacking in a cell-free system. IV. INTERACTION BETWEEN PRION PROTEINS According to the protein-only hypothesis, proposed by Prusiner (Fig. 5), the interaction of the cellular prion protein with the pathological isoform seems to be the crucial step in the conversion of PrPc to PrPSc. The existence of the hypothetical PrPc/PrPSc heterodimer may require the presence of a homodimer consisting of two PrPc molecules. This homodimer is thought to be in equilibrium with the PrPc monomers. It is unclear to date wether the spontaneous conversion reaction involves PrPc monomers or the PrPc homodimers.

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FIG. 5. Scheme of the conversion process of PrPC to PrPSc. Three possibilities for the conversion of PrPC into PrPSc do exist. An exogenous PrPSc triggers the conversion of PrP monomers leading directly to the hypothesized heterodimer consisiting of PrPC and PrPSc. Genetic predisposition of an individual leads to a spontaneous conversion of PrPC to PrPSc. The conversion process might proceed after formation of a dimeric PrPC or might occur with a monomeric PrPC. The central PrPSc heterodimer forms a PrPSc homodimer aggregating into amyloid fibrils.

In 1986 a 54 kDa protein was identified under denaturing conditions that may act as a dimeric PrP precursor for the scrapie protein (Bendheim and Bolton, 1986). A 60 kDa form of a recombinant hamster prion protein was detected in murine neuroblastoma cells in 1995 (Priola et al.,

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1995). It appears as a dimer under denaturing conditions analyzed by SDS-PAGE and under native conditions analyzed by immunoprecipitation. The linkage of both prion proteins might occur via hydrogen bonding, electrostatic interactions or covalent linkage involving lysins at the N-terminus of the protein. The observed dimer formation might be due to the hyperexpression of PrP with high PrP concentrations. The multimer formation of the prion protein and structural changes during this process has been investigated by fluorescence correlation spectroscopy (FCS) (Post et al., 1998). Prion aggregates mainly constituted of PrP27–30 were converted by sonication to monomeric PrP with an high α-helical content in the presence of 0.2% SDS. The oligomerization process was then initiated by the reduction of the SDS-concentration. Formation of β-sheet structured dimers was the initial step followed by oligomerization of these dimers within 10 minutes. After 1 hour PrP was aggregated. Whether the conversion reaction arises before the dimerization event or whether dimerization represents the initial step of the conversion process remains speculative. Prion proteins with mutations in the octarepeat region causing familial CJD show abnormal aggregation properties (Priola and Chesebro, 1998). Hamster PrPs encompassing two, four, and six octarepeats were expressed in mouse neuroblastoma cells. The fact that PrP dimers were detectable even under harsh denaturing conditions present in SDS-gel electrophoresis suggests that the PrP monomers were covalently linked rather than stabilized by noncovalent linkages such as hydrophobic interactions. However, covalently linked PrP dimers have still to be confirmed by other systems. Because of the lack of convincing experimental data, only a few models describe the PrP-dimerization process. One of them proposes the highly conserved region from aa 109 to aa122 as a major dimerization domain (Warwicker and Gane, 1996) calculated by a computational search for potential PrP interaction interfaces. Mutations such as alanine to valine at position 117 of human PrP associated with GerstmannSträussler-Scheinker syndrome reside within this region and might alter the stability of the dimer, facilitating the conversion of PrPc to PrPSc. In addition to the dimerization process, the association of the prion protein to the membrane could play an important role in TSE pathogenesis (Warwicker, 1999). The putative membrane-binding domain might be the first α helix. The agglomeration of the prion protein on the membrane might influence the orientation and configuration of PrP facilitating the PrP interaction process. Whether PrP dimers that have also been observed by us (Hundt, Gauczynski, Riley, and Weiss, manuscript in preparation) might play an

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important role in the PrP oligo/multimerization process and whether PrP/PrP interfering agents might hamper the entire PrP aggregation process have still to be investigated. V. OTHER PrP INTERACTING MOLECULES This section first describes PrP interacting molecules identified by ligand blots, yeast two-hybrid techniques or in vitro selection. Members of the PrP ligand family Pli are described followed by Bcl-2 belonging to the family of propoptotic and antiapoptotic molecules. Second, molecules are summarized acting as therapeutics in TSEs. With the exception of the protein clusterin, all the other molecules are of nonproteinaceous origin including polyanions, Congo red, polyene antibiotics, IDX, porphorins, and phtalocyanes. Finally, nucleic acids such as RNA aptamers are described in their function as PrP-interacting molecules. A. PrP Ligands (PLi’s) 1. Pli 45 and Pli 110 Two PrP binding proteins were identified in 1990, using ligand blots (Oesch et al., 1990). These two proteins identified from hamster brain were termed PrP ligands Pli 45 and Pli 110. To investigate the interaction of purified PrP with other proteins the authors used radiolabeled PrP27–30 and PrPc, respectively, for the binding of proteins from hamster brain that were separated by SDS-PAGE and blotted to nitrocellulose (ligand blots). Two major bands became visible by autoradiography using purified PrP27–30 and immunopurified PrPc. The molecular weight of the identified proteins were 45,000 and 110,000, respectively, and both proteins bound to PrPSc and PrPc derived from hamster brain. Other PrP-binding proteins ranging from 32–200 kDa were also observed. The stability of the complexes formed by Pli 45 and PrP 27–30 on nitrocellulose were investigated by intense washing steps and 50% of the radiolabeled PrP27–30 was washed off after 60 hours, corresponding to a dissociation rate constant of kD=3×10–6 s–1. Pli 45 revealed a sequence homology of 94.6% to murine GFAP (glial fibrillary acidic protein) at the cDNA level, suggesting that Pli 45 and GFAP are the same proteins. Comparative immunochemistry studies, using polyclonal Pli45- and GFAP-specific antibodies revealed the same staining pattern as monoclonal anti-GFAP antibodies in scrapie-infected sheep brain. In addition, both antibodies recognized recombinant

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GFAP expressed in Escherichia coli, suggesting that Pli 45 and GFAP are indeed the same proteins. Pli 45 was found exclusively in brain, whereas Pli 110 is present in several tissues, such as brain, lung, liver, spleen, and pancreas. Pli 110 was shown to be identical to PTP-associated splicing factor (PSF) (Oesch, 1994). Because studies with GFAPO/O mice revealed that GFAP is not essential for scrapie development (Gomi et al., 1995; Tatzelt et al., 1996a) and PSF is an essential splicing factor located in the nucleus (Patton et al., 1993), it seems that Pli45 and Pli110 do not play a crucial role in prion diseases. 2. Pli3-Pli8 Seven years after the identification of the first two PrP-binding proteins Pli 45 and Pli 110, six other PrP ligands were found (Table I) (Yehiely et al., 1997). The authors used a different system than that used for the identification of Pli 45 and Pli 110. Here, PrP was designed as a fusion protein with alkaline phosphatase (AP) and secreted by NIH 3T3 cells. PrP-AP was then used as a probe for screening the mouse brain cDNA library λgt11. Sequence analysis of nine clones revealed the six unique sequences, Pli3 to Pli8. Two cDNA clones showed homology to known sequences, to the mouse amyloid precursor-like protein (Aplp1) denoted Pli6 and to the mouse p45 NF-E2 related factor 2 (Nrf2), termed Pli7. All six Plis revealed the consensus sequence GXXXXXX(E/P)XP, which is not unique to PrP-binding proteins, but was identified in many other protein sequences. Hence, the authors concluded that it might represent a functional motif. Negative charge might also play a role in PrP binding, as four cDNA clones showed an excess of glutamic acids and aspartic acids over lysines and arginines. Each cDNA clone identified a single copy gene and the chromosomal location of each clone was identified in this work. Polyclonal antibodies directed against the polypeptides Pli3 and Pli5 were generated and purified. Both antibodies recognized proteins from N2a cells and mouse brain on Western blots. Anti-Pli3 antiserum detected a 70 and a 100 kDa polypeptide, whereas anti-Pli5 antiserum detected a 45 kDa polypeptide. All three identified polypeptides were believed to be novel PrP-binding proteins. Antisera to Nrf2 (Pli7) and Aplp1 (Pli6) were also used as probes on N2a cell lysates and mouse brain homogenates. For anti-Nrf2 antiserum, a 66 kDa protein was found that corresponds to the predicted size of mouse Nrf2. Aplp1 antiserum recognized polypeptides of about 85 and 95 kDa molecular mass, which are likely to be two different forms of Aplp1. The protein levels of Pli3 and Pli5 appeared similar in scrapie-infected and noninfected

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brain and N2a cells, whereas higher levels of Pli5 mRNA could be found in ScN2a cells. The protein levels of Nrf2 were found to be slightly decreased in ScN2a cells, whereas Aplp 1 protein levels remained unchanged in ScN2a cells and infected mouse brain. Higher mRNA levels for both Aplp1 and Pli5 were found in ScN2a cells. Aplp2 is a member of the APP-like (amyloid precursor protein) family, playing an important role in the pathogenesis of Alzheimer disease (AD). The major component of the senile plaques observed in AD is the Aβ peptide, which is derived from the APP protein (Glenner and Wong, 1984; Masters et al., 1985). PrP and Aplp1 are both membrane proteins; hence it is likely that they could interact on the cell surface. B. Bcl-2 Bcl-2 (Table I) represents a well-known member of a rapidly enlarging protein family of proapoptotic and antiapoptotic molecules, including at least 15 related proteins (Adams and Cory, 1998). In 1995 the role of Bcl-2 was investigated using a yeast two-hybrid screen (Kurschner and Morgan, 1995). LexA-Bcl-2 in the bait and a murine cerebellar cDNA-VP16 fusion library in the prey position identified potential Bcl-2 binding proteins. Surprisingly, the prion protein and not bax, which is known to heterodimerize with Bcl-2 (Oltvai et al., 1993), was pulled out by this screen. The sequenced cDNA clone contained a fusion between the VP16 domain and mouse PrP, encompassing aa72 to aa245, denoted PrP-VP16. Using LexA-PrP in the bait and Bcl-2-VP16 in the prey position of the yeast two-hybrid system resulted also in an interaction between PrP and Bcl-2. Interactions with other members of the Bcl-2 family, such as Bax or A1 were not observed. The PrP mutation P102L, associated with human Gerstmann-SträuslerScheinker syndrome was investigated, and it was shown that this mutation did not alter the binding behavior of PrP to Bcl-2. Interestingly, the PrP-Bcl-2 interaction could not be confirmed by coimmunoprecipitation assays, suggesting that this protein interaction can be observed only in the yeast-two-hybrid system. Bcl-2 and Bax act as antiapoptotic and proapoptotic molecules in apoptosis, respectively. Moreover, the ratio of Bax-Bcl-2 heterodimers to homodimers of each protein is important for the regulation of apoptosis (Oltvai and Korsmeyer, 1994; O’Dowd et al., 1988; Yang and Korsmeyer, 1996). Hence the authors concluded that PrP might play a role in disrupting the Bax:Bcl-2 ratio by trapping Bcl-2 and favoring Bax-Bax homodimers, which would lead to cell death by apoptosis (Fig. 6). The trapping of Bcl-2 by PrP might occur during trafficking of PrP before

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FIG. 6. Schematic view of Bcl-2::Bax, Bcl-2::PrP, and Bax::Bax transitions and their possible role in cellular functions.

exposure to the cell membrane. Although Bcl-2 and PrP are both membrane associated, the physiological cellular location of Bcl-2 is different from that of PrP. Bcl-2 is thought to be an inner mitochondrial membrane protein (Hockenbery et al., 1990; Motoyama et al., 1998) or might reside on the mitochondrial outer membrane, the endoplasmatic reticulum, or the nuclear membrane (Krajewski et al., 1993; Lithgow et al., 1994), and is not present on the cell surface membrane. C. Laminin Laminin (LN) is a glycoprotein of the extracellular matrix (ECM) [for review see Beck et al. (1990)] that mediates cell attachment, communication, differentiation, movement, and neurite outgrowth promotion (Hunter et al., 1989). Laminin is the first ECM protein detected during embryogenesis. In later development and in mature tissue, laminin serves as a ubiquitious and major noncollagenous component of basement membranes (Beck et al., 1990). Laminin was first isolated from Engelbreth-Holm-Swarm (EHS) tumor (Timpl et al., 1979) and from extracellular deposits of murine parietal yolk sac (PYS) carcinoma cells (Chung et al., 1979). A specific binding between laminin and the amyloid precursor protein (APP), the precursor of the amyloid peptide involved in Alzheimer’s disease, has been identified (Narindrasorasak et al., 1992). APP and β-amyloid peptide (1–40) interaction with the extracellular matrix promotes neurite outgrowth, suggesting that the complex might play a normal physiological role in the brain (Kibbey et al., 1993; Koo et al., 1993). Recently, a direct interaction between the cellular prion protein (PrPc) and laminin was reported (Graner et al.,

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2000). An involvement of the PrPc-laminin interaction in neuritogenesis induced by NGF plus laminin in the PC-12 cell line was further suggested (Graner et al., 2000). Neuritogenesis, induced either by laminin or its γ-1-derived peptide in primary cultures from rat or either wildtype or PrP null mice hippocampal neurons, might imply that PrPc could be the main cellular receptor for the particular γ-1 domain located in the carboxy terminus of laminin (Graner et al., 2000). D. Therapeutics 1. Polyanions Polyanions (Table IV), including heteropolyanion 23 (HPA-23), Dextran Sulfate 500 (DS 500), pentosan polysulfate (SP54), and heparin are known to bind the prion protein and/or prevent PrPSc accumulation in animals and cell systems (Brimacombe et al., 1999; Caughey and Raymond, 1993; Diringer and Ehlers, 1991; Ehlers and Diringer, 1984; Farquhar et al., 1999; Gabizon et al., 1993; Kimberlin and Walker, 1983; Kimberlin and Walker, 1986; Ladogana et al., 1992). The first polyanion denoted as an anti-scrapie drug was HPA-23 (Kimberlin and Walker, 1983; Kimberlin and Walker, 1986). The effect of HPA-23 was tested in several different scrapie strains, such as 139A, ME7, 22A, and 263K. HPA-23 was effective in all these strains and prolonged the lifetimes of the animals significantly after scrapie injection. Less effect was observed when scrapie material was injected intraperitoneally or if the drug was given more than 48 hours after scrapie infection. Injection before infection with scrapie is not effictive, owing to the rapid metabolization or excretion of HPA-23. HPA-23 is thought to interfere with early replication of PrPSc in the lymphoreticular system, reducing the efficiency of scrapie infection. These results, together with the brain toxicity of this molecule, suggest that HPA-23 has limited therapeutic value. Two high-molecular-weight polyanions, carrageenan and DS 500, were shown to be highly efficient in reducing scrapie titers in mice infected with the 139A strain of scrapie (Ehlers and Diringer, 1984; Kimberlin and Walker, 1986). All intravenous or intraperitoneal combinations of injecting DS 500 or scrapie reduced the effective titer about 100- to 200-fold. The effect of DS 500 is long-lasting. Application of DS 500 up to 10 weeks before infection increases the incubation period in mice. However, DS 500 itself is highly toxic and causes up to 50% mortality at a dose of 2 mg per mouse. Like HPA-23, DS 500 is thought to prevent PrPSc replication in spleen and lymph nodes, and its mode of action is likely to be independent of its activity as a B-cell mitogen. The

256 TABLE IV Antiscrapie Drugs Likely to Interact Directly with PrP Tested Scrapie Strain

Drug

Successfully Treated Animals

HPA-23

139A, ME7, 22A and 263K

Mouse and hamster

DS 500

139A

Mouse

Pentosan Polysulfate

139A, ME7, 22A, and 263K

Mouse and hamster

Suggested Mode of Action

Comments

References

Prevents early agent Effective in a lot of scrapie Kimberlin and Walker replication in the LRS, strains, rapid metabolism (1983); Kimberlin and competes with GAG and excretion, toxic Walker (1986) (glycosaminoglycan) binding site Prevents agent replication Long-lasting antiscrapie Ehlers and Diringer (1984); in the LRS due to its high effect but toxic at Kimberlin and Walker molecular weight and therapeutic doses (1986) negative charge, competes with GAG (glycosaminoglycan) binding site Interferes with PrPSc uptake Very promising drug, Ehlers and Diringer (1984); from nerve endings, effective at extreme low Farquhar et al. (1999); competes with GAG dose Ladogana et al. (1992) (glycosaminoglycan) binding site

Amphotericin B

C506M3 and 263K

Mouse and hamster

MS-8209

C506M3 and 263K

Mouse and hamster

Congo red

263K and 139A

Hamster

Anthracycline

263K

Hamster

Porphyrins and Phtalocyans

236K

Cp-60/Cp-62

ScN2a cells

Mouse expressing Hamster PrP None

iPrP13 (b-sheet breaker) Clusterin [apolipoprotein J (apo J)]

139A

Mouse

None, prevents aggregation of PrP106–126

Binding to PrPc/Sc

Direct prevention of PrP Acute nephrotoxicity and Pocchiari et al. (1987); Xi et conversion or interference low solubility, widely used al. (1992) with PrPSc uptake for the treatment of fungals Same as for AmB Lower toxicity than AmB Adjou et al. (1995); Demaimay et al. (1997); Adjou et al. (1999) Binding to PrPc with Dyes amyloid Caspi et al. (1998); Caughey polyanion-like behavior, et al. (1993); Ingrosso et or binding to PrPSc al. (1995) (overstabilisation) Binding to PrPSc, preventing Used for the treatment of Tagliavini et al. (1997) amyloid deposition malignancies Binding to PrPc/Sc Inhibits cell free PrPc/Sc Caughey et al. (1998); Priola conversion et al. (2000) Mimicking donimant negative inhibition of prion replication Direct change of PrP secondary structure Binds to extraneuronal PrPBSE

Identified by using a computational database search Synthetic peptide

Perrier et al. (2000)

Soto et al. (2000) McHattie and Edington (1999)

257

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high-molecular-weight and negative charge may represent important factors in the antiscrapie effect of DS 500. SP54 (Pentosan Polysulfate, Fig. 7A) has an antiscrapie effect comparable to DS 500, but is less toxic. It has been shown that SP54 significally increases scrapie incubation period in hamsters infected with 263K scrapie strain and in mice infected with the 139A, Me7 and 22A strains of scrapie (Ehlers and Diringer, 1984; Farquhar et al., 1999; Ladogana et al., 1992). SP54 is even effective if only a single low-dose is injected after infection. A single injection of 250 µg of SP54 increased the mean incubation period of the ME7 strain by up to 66% and 1 mg of SP54 protected mice completely from the 22A scrapie strain. SP54 is thought be effective during the very early events of pathogenesis by interfering with the uptake of PrPSc by nerve endings and/or carrier cells. The low-dose effect and the lower in vivo toxicity compared to other polyanions make SP54 a promising candidate in the field of antiscrapie polyanions. All antiscrapie polyanions published so far might act by competing directly with the binding of cellular glycoaminoglycans (GAGs) to PrPC (see section II. A) and/or PrPSc (Brimacombe et al., 1999; Caughey et al., 1994). Indeed, GAGs are involved in the metabolism of PrPc (see section II.A) and thus in the biogenesis of PrPSc. It was shown by surface plasmon resonance, that pentosan polysulfate shows the strongest binding to recombinant PrP followed by heparin and dermatan sulfate. This correlates to the ability of the molecules to delay scrapie disease and reduce PrPSc accumulation in scrapie-infected cell lines (Caughey and Raymond, 1993). 2. Congo Red Congo red (Fig. 7C, Table IV) is a dye that can be used as a diagnosic stain for amyloids. It is well known that Congo red can inhibit PrPres accumulation in Sc+-MNB cells and PrPSc replication in 263K and 139H treated hamsters (Caspi et al., 1998; Caughey et al., 1994; Caughey et al., 1993; Ingrosso et al., 1995). The mechanism of the Congo red antiscrapie effect probably involves direct binding to PrPc, which again is thought to block the binding of cellular GAGs to PrPc, as described for polyanions (Caughey et al., 1994). The proposed direct binding of Congo red to PrPSc is thought to stabilize PrPSc, the abnormal isoform of the prion protein, and prevents its partial denaturation, which could be necessary for agent replication (Caspi et al., 1998). 3. Polyene Antibiotics Amphotericin B (AmB) and MS-8209 (Fig. 7D) are polyene macrolide antibiotics that have a ring structure containing a hydrophobic and a

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FIG. 7. Antiscrapie drugs of four different classes. (A) Pentosan polysulfate as a powerful drug belonging to the polyanion family. (B) IDX a derivative of doxorubicin. (C) Congo red belonging to the diazo dyes. (D) Amphotericin B and MS-8209 belonging to the family of polyene macrolide antibiotics.

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TABLE V Antiscrapie Drugs not Thought to Interact Directly with PrP

Drug

Tested on Scrapie Strain

Success in Animal Treatment

Dapsone

SY

Mouse

Flurpirtine (Katadolon)

—

None, cures neuronal cells treated with PrP106–126

Suggested Mode of Action

References

Altering of macrophage Manuelidis processing of infectious et al. (1998) agent and modulation of inflammatory factors Lowers toxic effect of Perovic et al. PrP106–126 by (1995) normalization of GSH levels

hydrophilic region on either side of the molecule. They are used for the treatment of systemic fungal infections such as candidiasis, histoplasmosis, and aspergillosis (Medoff et al., 1983). The effects of AmB and its derivative MS-8209 were studied in several models of rodents including 263Kinfected hamsters. Both were very efficient in delaying scrapie disease and PrPSc accumulation. MS-8209 shows at least a five times lower toxicity and a higher solubility and is able to double the incubation time of scrapie in hamsters. In contrast to polyanions, polyene antibiotics are effective even after intracerebral infection (Adjou et al., 1995; Demaimay et al., 1994; McKenzie et al., 1994; Pocchiari et al., 1987; Xi et al., 1992). Presently AmB and its derivatives are the only category of antiscrapie drugs that are prolonging the incubation period when given at late stages of infection (Demaimay et al., 1997). However, the effect of polyene antibiotics vary between scrapie strains (Adjou et al., 1996). Note that the only reported treatment of clinical CJD with AmB in humans was unsuccessful (Masullo et al., 1992). Several possible mechanisms are involved in the antiscrapie effect of polyene antibiotics. AmB and MS-8209 have been proposed to directly affect the PrPsen to PrPres conversion step and thus prevent PrPres accumulation (Adjou et al., 1999; Adjou et al., 1997; Demaimay et al., 1997). Nevertheless a more indirect mode of action seems to be possible, whereby AmB and its derivatives disturb the uptake of PrPres by cells most likely by interfering with membrane cholesterol-rich domains (rafts) (Bolard, 1986; Taraboulos et al., 1995). 4. Other Therapeutics Anthracycline 4′-iodo-4′-deoxy-doxorubicin (IDX) (Fig. 7B; Table IV) is a derivative of the drug doxorubicin, which is successfully used

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in the treatment of several malignancies (Barbieri et al., 1987). IDX binds to amyloid fibrils and induces amyloid resorption in patients suffering from plasma cell dyscrasias with immunoglobulin lightchain amyloidosis (Gianni et al., 1995; Merlini et al., 1995). IDX was shown to delay the clinical signs of scrapie disease in 263K-infected hamsters when co-incubated with the 263K material before to intracerebral inoculation. At a molecular level IDX is thought to bind the abnormal form of PrP, thereby decreasing the number of template molecules available for the PrPc conversion process (Tagliavini et al., 1997). Porphyrins and phtalocyans (Table IV) prevented PrPres accumulation in scrapie-infected mouse neuroblastoma cell cultures (Caughey et al., 1998) and prolonged the incubation period in hamster PrP expressing mice infected with 263K scrapie (Priola et al., 2000). The molecules also inhibited a cell-free conversion of hamster PrPsen to PrPres, showing that the effect seems to be due to direct PrP-binding. Nevertheless, because PrPres preparations are not completely pure, interactions with other molecules might be possible. Some other interactions with cells involved in scrapie pathogenesis can also not be excluded (Manuelidis, 2000). Based on the proposal of a protein X binding domain (Kaneko et al., 1997b) synthetic drugs were identified that are able to inhibit PrPSc formation in ScN2a cells (Perrier et al., 2000). Two compounds, Cp-60 and Cp-62 (Table IV), act in a dose-dependent manner and show low toxicity. They are suggested to mimic the dominant negative inhibition of PrP replication originally reported for a PrP mutant (Kaneko et al., 1997a). A 13-residue β-sheet breaker peptide (iPrP13) (Table IV) was shown to partly reverse PrPSc to a PrPc-like state. Mice inoculated with iPrP12pretreated infectious material showed delayed appearance of clinical symptoms (Soto et al., 2000). The peptide is thought to directly change the conformation of PrPSc from a β-sheeted to a more α-helical secondary structure and therefore reduce infectivity. An effect of clusterin (Table IV) on the in vitro aggregation of the prion neuropeptide 106–126 was tested. Clusterin co-localizes with extraneuronal PrPBSE in terminal BSE and the aggregation of the neuropeptide 106–126 was inhibited by clusterin in a dose-dependent manner (McHattie and Edington, 1999). The neurotoxicty of peptide 106–126 is a subject of discussion, as a recent report described aggregation but no neurotoxicity for this peptide (Kunz et al., 1999). Dapsone (Manuelidis et al., 1998) and flurpirtine (Perovic et al., 1995) have also been described as TSE therapeutics. In contrast to the previously described drugs, however, a direct interaction with PrP here is unlikely (Table V).

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E. Nucleic Acids So far, no nucleic acid directly linked to scrapie infectivity has been identified. The existence of scrapie-specific homogeneous nucleic acid of more than 80 nucleotides has been excluded by analysis of highly purified scrapie preparations involving improved return refocusing gel electrophoresis (Kellings et al., 1992). However, the presence of a nucleic acid associated with infectivity cannot be ruled out, as the BSE agent can be transmitted to mice in the absence of detectable abnormal PrP (Lasmézas et al., 1997). The in vitro interaction of nucleic acid with PrP has been described for both DNA and RNA. Using fluorescence labeled DNA, it was shown that the binding strength of peptide PrP106–126 to DNA was of a similar order of magnitude as the binding of retroviral protein p10 with model nucleic acids (Nandi, 1997). It was also shown that PrP106–126 polymerizes in the presence of DNA in solution, whereas the peptide alone fail to polymerize (Nandi, 1998). RNA aptamers that bind specifically to recombinant hamster PrP (Weiss et al., 1995) but not to recombinant PrP90–231 (Weiss et al., 1996) were isolated by in vitro selection (Weiss et al., 1997). RNA aptamers of three different motifs were isolated, and all revealed a G quartet scaffold, which was proved to be essential for PrPc binding. An RNA aptamer of only 29 nucleotides, representing the G quartet scaffold, was sufficient for PrPc recognition. The interaction of the G quartet scaffold with PrPc was directed exclusively against the amino terminus (aa23–52) of PrP. However, it could not be excluded that the aptamer recognizes PrPSc, but failed to recognize PrP27–30, lacking aa23–89 from the amino terminus. ACKNOWLEDGMENTS We thank the Bundesministerium für Bildung, Forschung, Wissenschaft und Technologie (BMBF) grant # KI 01-9760 and the European Union (EU) grants # FAIR-5-CT97-3314, #FAIR6-CT98-7020 and #BM-H4-98-6054 for financial support. We thank Corinne Ida Lasmézas and Louise Riley for critical reading of the manuscript. S.W. thanks Rudolf Grosschedl and Ernst-Ludwig Winnacker for valuable advice and continuous support.

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Tatzelt, J., Maeda, N., Pekny, M., Yang, S.L., Betsholtz, C., Eliasson, C., Cayetano, J., Camerino, A.P., DeArmond, S.J., and Prusiner, S.B. (1996a). Scrapie in mice deficient in apolipoprotein E or glial fibrillary acidic protein. Neurology 47, 449–453. Tatzelt, J., Prusiner, S.B. and Welch, W.J. (1996b). Chemical chaperones interfere with the formation of scrapie prion protein. EMBO J. 15, 6363–6373. Tatzelt, J., Zuo, J., Voellmy, R., Scott, M., Hartl, U., Prusiner, S.B., and Welch, W.J. (1995). Scrapie prions selectively modify the stress response in neuroblastoma cells. Proc. Natl. Acad. Sci. U.S.A. 92, 2944–2948. Telling, G.C., Scott, M., Mastrianni, J., Gabizon, R., Torchia, M., Cohen, F.E., DeArmond, S.J., and Prusiner, S.B. (1995). Prion propagation in mice expressing human and chimeric PrP transgenes implicates the interaction of cellular PrP with another protein. Cell 83, 79–90. Timpl, R., Rohde, H., Robey, P.G., Rennard, S.I., Foidart, J.M., and Martin, G.R. (1979). Laminin-a glycoprotein from basement membranes. J. Biol. Chem. 254, 9933–9937. Tobler, I., Gaus, S.E., Deboer, T., Achermann, P., Fischer, M., Rülicke, T., Moser, M., Oesch, B., McBride, P., and Manson, J.C. (1996). Altered circadian activity rhythms and sleep in mice devoid of prion protein. Nature 380, 639–642. Vey, M., Pilkuhn, S., Wille, H., Nixon, R., DeArmond, S.J., Smart, E.J., Anderson, R.G., Taraboulos, A., and Prusiner, S.B. (1996). Subcellular colocalization of the cellular and scrapie prion proteins in caveolae-like membranous domains. Proc. Natl. Acad. Sci. U.S.A. 93, 14945–14949. Wang, K.S., Kuhn, R.J., Strauss, E.G., Ou, S., and Strauss, J.H. (1992). High-affinity laminin receptor is a receptor for Sindbis virus in mammalian cells. J. Virol. 66, 4992–5001. Warwicker, J. (1999). Modelling charge interactions in the prion protein: predictions for pathogenesis. FEBS. Lett. 450, 144–148. Warwicker, J., and Gane, P.J. (1996). A model for prion protein dimerisation based on alpha-helical packing. Biochem. Biophys. Res. Commun. 226, 777–782. Weiss, S., Famulok, M., Edenhofer, F., Wang, Y.H., Jones, I.M., Groschup, M., and Winnacker, E.L. (1995). Overexpression of active Syrian golden hamster prion protein PrPc as a glutathione S-transferase fusion in heterologous systems. J. Virol. 69, 4776–4783. Weiss, S., Proske, D., Neumann, M., Groschup, M.H., Kretzschmar, H.A., Famulok, M., and Winnacker, E.L. (1997). RNA aptamers specifically interact with the prion protein PrP. J. Virol. 71, 8790–8797. Weiss, S., Rieger, R., Edenhofer, F., Fisch, E., and Winnacker, E.-L. (1996). Recombinant prion protein rPrP27–30 from Syrian Golden Hamster reveals proteinase K sensitivity. Biochem. Biophys. Res. Commun. 219, 173–179. Weissmann, C. (1996). PrP effects clarified. Curr. Biol. 6, 1359. Welch, W.J., and Brown, C.R. (1996). Influence of molecular and chemical chaperones on protein folding [published erratum appears in Cell Stress Chaperones 1996 Sep; 1(3):207]. Cell Stress Chaperones 1, 109–115. Wewer, U.M., Liotta, L., Jaye, M., Ricca, G.A., Drohan, W.N., Claysmith, A.P., Rao, C.N., Wirth, P., Coligan, J.E., Albrechtsen, R., Mudry, M., and Sobel, M.E. (1986). Altered levels of laminin receptor mRNA in various human carcinoma cells that have different abilities to bind laminin. Proc. Natl. Acad. Sci. U.S.A. 83, 7137–7141. Xi, Y.G., Ingrosso, L., Ladogana, A., Masullo, C., and Pocchiari, M. (1992). Amphotericin B treatment dissociates in vivo replication of the scrapie agent from PrP accumulation [see comments]. Nature 356, 598–601.

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TRANSGENIC STUDIES OF THE INFLUENCE OF THE PrP STRUCTURE ON TSE DISEASES BY EMMANUEL A. ASANTE AND JOHN COLLINGE MRC Prion Unit and Department of Neurogenetics, Imperial College School of Medicine at St. Mary’s, London W2 1PG, United Kingdom

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Effects of PrP Gene Ablation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. PrPC is Necessary for Disease Propagation . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Neurografting of PrPC Producing Cells in PrP Null Mice . . . . . . . . . . . . B. Conditional Knockout of Prnp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Structure and Function of the PrP Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. N-Terminal Deletions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Internal Deletions of PrP Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Genetic Link of PrP Gene to Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Transgenic Studies of PrP Topology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. Spontaneous Disease in Mutant Transgenic Mice . . . . . . . . . . . . . . . . . . . . . VII. Transgenic Studies of the Species Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The Concept of Species Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Murine to Hamster Species Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Artificial Prions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Human to Murine Species Barrier. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Bovine to Murine Species Barrier. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Bovine to Human Species Barrier. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Transgenic Studies of Incubation Period . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX. Transgenic Studies of the Molecular Basis of Prion Strains . . . . . . . . . . . . . . A. What are Strains? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Molecular Strain Types . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. What is the Cause of Neurodegeneration in Prion Diseases?. . . . . . . . . . X. Transgene Vector Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . XI. Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. INTRODUCTION Prion diseases are a group of rare transmissible neurodegenerative disorders that affect a range of mammalian species including humans. The human prion diseases have traditionally been classified into Creutzfeldt-Jakob disease (CJD), Gerstmann-Sträussler-Scheinker disease (GSS), and kuru. With a worldwide incidence of ~1 person per million, these rare diseases have recently attracted remarkable attention because of both their unique biology and concerns that the epidemic of the newly recognized bovine spongiform encephalopathy 273 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

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(BSE) could pose a threat to public health through dietary exposure to infected tissues. The transmissible agent, or prion, seems to consist principally of an abnormal isoform of the prion protein (PrP), designated PrPSc. PrPSc is known to be derived from the cellular isoform, PrPC, by a posttranslational mechanism and evidence is mounting that this change is conformational rather than covalent. Although PrPC is fully sensitive to proteolysis, PrPSc, which accumulates in the brain during disease, is partially protease resistant. PrPC is predominantly α helical and is fully sensitive to proteolysis, whereas PrPSc has a high β-sheet content and is partially protease resistant (Caughey et al., 1991; Pan et al., 1993). The human PrP gene (PRNP) is a single copy gene located on the short arm of chromosome 20; it spans 16 kb and contains two exons. The complete open reading frame of 759 nucleotides is contained within the larger second exon, which accounts for the majority of the 2.4 kb mRNA. There are doubts about the structural characteristics of the infectious form of PrP; however, it is well established that prion diseases arise by one of three processes. In outline all three etiological routes can be described with reference to a single, general model. In this model the native PrPC molecule is in equilibrium with the rare PrPSc conformational isoform. PrPSc can then be stabilized by complementary association with a like molecule or can actively convert PrP chains to a like conformation. Assembly then continues until a stable seed is formed. Such structures can continue to grow by accretion and can divide by breakage into smaller, infectious units. A mechanism involving linear polymerization has been modeled mathematically and predicts many of the observed aspects of the disease (Masel et al., 1999). This gross mechanism explains the observation that prion diseases occur by inherited mutations (Hsiao et al., 1989; Collinge, 1997), which destabilize the cellular form and therefore predispose it to conversion to PrPSc or by iatrogenic or dietary infection with PrPSc. Sporadic cases are also described where the cause is unknown but within the preceding paradigm they can be explained either by somatic mutation or by a rare, spontaneous conversion of the wild-type protein to the PrPSc conformation. Cloning and characterization of the mouse (Basler et al., 1986), human (Kretzschmar et al., 1986), and the hamster (Scott et al., 1989) prion protein genes paved the way for transgenic studies to contribute a great deal of new knowledge to our understanding of prion biology. The existence of inherited mutations and the practical reality of introducing foreign genes into the germ line of mammals allow the creation of animal models designed to answer specific questions on prion biology. Transgenic technology has been used extensively to study the struc-

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ture and function of the prion protein and its influence on disease propagation and progression. In this chapter we review the advances in prion biology facilitated by transgenic technology, with emphasis on structure-function relationships of the prion gene. II. EFFECTS OF PrP GENE ABLATION A major prediction from the protein-only hypothesis is that mice devoid of PrPC will lack the template for conversion to PrPSc and should therefore be resistant to experimental prions. In 1992 Büeler and others reported on the first PrP knockout mice (Zurich I), which, contrary to expectation for a gene that is active during embryonic development (Manson et al., 1992), exhibited normal development and behavior (Büeler et al., 1992). A second PrP knockout line was independently created (Manson et al., 1994) that corroborated the findings of Büeler and co-workers. However, electrophysiological studies have demonstrated impaired GABAA (γ-aminobutyric acid type-A) receptor-mediated synaptic inhibition in hippocampal brain slices maintained in vitro, as well as impaired long-term potentiation in these two initial knockout lines (Collinge et al., 1994; Manson, et al., 1995). These abnormalities of synaptic inhibition are reminiscent of the neurophysiological abnormalities seen in patients with CJD and in scrapie-infected mice (Jefferys et al., 1994). This raises the possibility that prion neurodegeneration may be, at least in part, due to loss of PrP function rather than to a deleterious effect of PrPSc (Collinge et al., 1994). Two groups, using different methods, notably smaller inhibitory postsynaptic currents (IPSCs) and subphysiological temperatures, failed to replicate this result in cerebellar (Herms et al., 1995) and hippocampal (Lledo et al., 1996) slices. Further, neurophysiological abnormalities, namely a reduction of slow afterhyperploarization (AHPs) evoked by trains of action potentials in the hippocampus of PrP null mice, including disrupted Ca2+activated K+ currents and abnormal intrinsic properties of CA1 pyramidal cells, have been reported (Colling et al., 1995; Colling et al., 1996). This PrP null phenotype has subsequently been confirmed by another group (Herms et al., 1998). Also an alteration in circadian rhythms, as well as disturbance in sleep patterns, has been described in these knockout mice (Tobler et al., 1996). A third PrP knockout line (Sakaguchi et al., 1996) posed a conundrum in that these mice developed a severe cerebellar ataxia and Purkinje cell degeneration at about 70 weeks of age. Subsequent to these three knockout lines, two more PrP-deficient lines of mice have been generated that

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also develop late-onset ataxia (Cozzio et al., 1999; Moore, 1997). The first notable difference between the ataxic and nonataxic PrP knockout lines lies in the strategy used in the gene inactivation process. For instance, in the nonataxic Zurich I line (Büeler et al., 1992), the PrP open reading frame was disrupted by replacing 184 of 254 codons with the neo resistance gene. Manson et al., (1994) inserted a selectable marker into a unique KpnI site of the PrP ORF without any significant deletions. However, the ataxic Nagasaki line (Sakaguchi et al., 1996) had deletion of the entire coding sequence, about 900 bp of intron 2 and about 450 bp of 3′ untranslated sequence. The Zurich group then addressed the issue of whether the ataxic phenotype was due to ablation of the prion gene per se or due to the loss of essential genetic information in the vicinity of the PrP coding sequence by generating the Zurich II knockout line (Cozzio et al., 1999). In this line approximately 250bp of intron 2, the entire PrPencoding exon 3 and about 600bp of 3′ untranslated sequence were deleted by Cre/LoxP mediated recombination (Cozzio et al., 1999). These mice with a more substantial deletion of PrP sequences showed a phenotype similar to the ataxic Nagasaki mice (Sakaguchi et al., 1996), presenting with abnormal gait, motor impairment of the hindlegs, and Purkinje cell degeneration (Cozzio et al., 1999). Similarly, Moore (1997) generated the Rcm0 PrP-deficient line by the strategy of double replacement in ES cells, resulting in the deletions of a significant amount of the PrP sequence and obtained an ataxic phenotype. An important question that remains unanswered is whether the ataxic phenotype is a direct result of PrP inactivation or indirect effect of other genes being activated or inactivated as a result of the gene targeting strategy. It is now known that the effects of a neighboring gene that is being inadvertently upregulated when as yet to be determined 5′ and 3′ PrP sequences are deleted may in some way explain the ataxic phenotype. Moore et al. (1999) detected an upregulation of a novel locus Prnd designated doppel (Dpl) in the central nervous system (CNS) of the Nagasaki and Rcm0 lines that develop late on-set ataxia and Purkinje cell degeneration but not in the Zurich I lines. The prnd is found 16 kb downstream of the mouse prion protein gene Prnp and encodes a 179 residue PrP-like protein with about 25% identity with all known prion proteins (Moore et al., 1999). Unlike the Prnp, the Prnd is expressed minimally in the adult CNS and appears to be upregulated in the CNS in PrP-deficient mice, which develop cerebellar ataxia. When the targeted alleles involve a disruption of PrP ORF, an invasion of intron 2 and removal of the 3′ splice acceptor site, then there is upregulation of Prnd in the brain. The upregulation of Prnd mRNA results from Prnp-Prnd intergenic splicing

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driven from the Prnp promoter rather than the Prnd promoter (Moore et al., 1999). The overexpression of Dpl may provoke neurodegeneration and cerebellar dysfunction seen in the Nagasaki and Rcm0 PrP knockout lines rather than PrP deficiency (Moore et al., 1999). It would be reasonable to predict that the Zurich II lines, which also exhibit ataxia and Purkinje cell degeneration, would have similar upregulation of Prnd in the brain. One of the standard methods of confirming the effect of knocking out a gene where a clear null phenotype results is to reintroduce a functional copy of the gene understudy into the knockout background. A complete restoration of the null phenotype would suggest the correct gene has been inactivated; therefore any observed phenotype could be ascribed to disruption of the function of the gene in question. However the method used for the complementation may be important. Ideally targeting of the disrupted locus by homologous recombination to replace it with a functional copy is to be preferred. This method would ensure that identical copy number and genetic environment are maintained, but this is not always possible or easy. Alternatively, the appropriate transgene could be microinjected into eggs of the knockout mice and transgenics generated. The advantage of these two approaches is that only expression of the disrupted gene is restored. However the microinjection approach could tip the balance the other way in cases where a possible overexpression can itself lead to a phenotype, though it appears not be a common problem. A third approach is to mate the knockout mice to wild-type mice, but this approach, although quick and easy, has a major drawback. In situations such as PrP knockouts where there is controversy and the possibility of other genes controlling the observed phenotype cannot be excluded, complementation by mating will give ambiguous results. Complementation experimentation has been done in the ataxic mice (Sakaguchi et al., 1996) by mating, instead of inserting only PrP transgene by microinjection into the eggs from the null mice (Nishida et al., 1999). Because the mating approach is nonselective, any unidentified gene whose inadvertent disruption might be causing the ataxic phenotype will be rescued as well. When the ataxic Sakaguchi knockout mice were crossed with transgenic mice haboring a wild-type mouse PrP-A transgene designated Tg(MoPrP-A)4053/FVB, none of the positive Tg(P)Prnp o/o presented with ataxia at more than 90 weeks of age (Nishida et al., 1999). The number and topography of Purkinje cells were preserved in all areas of the cerebellar cortex in the Tg(P)Prnp o/o mice. The authors concluded that the successful rescue of the ataxic phenotype in the Ngsk Prnp o/o

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argued against the effects of random mutations in genes other than the Prnp locus and that disruption of the Prnp gene was responsible for the Purkinje cell loss. Similarly, Büeler et al. (1993) introduced hamster PrP transgenes into the Zurich I knockout mice by mating to transgenic mice expressing hamster PrP (Scott et al., 1989) and rendered them susceptible to hamster but not mouse scrapie infection. Furthermore, the neurophysiological phenotype found in the Zurich I mice (Collinge et al., 1994) was rescued by introduction of normal PrP alleles through mating of the null mice with transgenic mice expressing wild-type human PrP (Whittington et al., 1995). The data of Cozzio et al., (1999) on a second knockout line by the Zurich group (Zurich II) in which intron 2 has been similarly removed appear to support the ataxic phenotype. However, it has now been suggested that the doppel gene, which is upregulated in the ataxic knockouts, may be responsible for the ataxia and Purkinje cell loss (Moore et al., 1999). On the basis of the successful complementation of the different PrP knockout phenotypes, it may be surmized that irrespective of the knockout strategy, the prion gene is being adequately inactivated, and that the ataxic phenotype and cerebellar Purkinje cell loss are artifacts of particular knockout strategies. If indeed PrPC has a function that is masked by the doppel gene, knocking out both doppel and PrP simultaneously in the mouse might give indications of the elusive function of the prion protein. III. PrPC IS NECESSARY FOR DISEASE PROPAGATION In spite of the differences in strategy and outcome, the most important revelation from all the knockout lines discussed above was that when challenged with scrapie they were completely resistant to disease and did not propagate infectivity (Büeler et al., 1993; Sailer et al., 1994; Manson et al., 1994; Sakaguchi et al., 1995). Moreover, mice heterozygous for PrP gene ablation showed prolonged incubation times (Büeler et al., 1994). This demonstrates unequivocally that PrPC is required for disease propagation and progression. A. Neurografting of PrPC Producing Cells in PrP Null Mice Further evidence of the central role of PrPC is provided by a series of neurografting experiments in PrP null mice. Aguzzi and co-workers engrafted PrP-overproducing embryonic neuroectodermal tissue from tga20 mice (Fischer et al., 1996) into the brains of Prnpo/o mice

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immunotolerant to PrP, and not susceptible to scrapie. They inoculated the engrafted mice with scrapie prions and observed that infected grafts developed severe histopathological changes characteristic of scrapie and contained high amounts of PrPSc and infectivity, while neighboring cells deficient of PrPC remained unaffected (Brandner et al., 1996a). A substantial amount of graft-derived PrPSc migrated into the host brain, but 16 months after inoculation no pathological changes were seen in PrP-deficient tissue, not even in the immediate vicinity of the grafts (Brandner et al., 1996a). The host life span was not reduced, leading to the conclusion that availability of endogenous PrPC to the infectious agent, rather than deposition of PrPSc, correlates with scrapie neurotoxicity in vivo. They then addressed the spread of prions from peripheral sites to the CNS by intraocular administration of scrapie to Prn-p o/o mice engrafted with PrPC overproducing neuroectoderm, but did not detect pathology or PrPSc in the grafts up to 66 weeks (Brandner et al., 1996b). However, scrapie prions inoculated intraocularly into wild-type mice spread efficiently to the CNS within 16 weeks. To rule out the possibility that prion transport was disabled by a neutralizing immune response, the experiments were repeated in transgenic mice overexpressing PrP on T lymphocytes under the control of the lck promoter (Raeber et al., 1999). These mice (designated tg33) were resistant to scrapie and did not contain scrapie infectivity in brain and spleen after inoculation with scrapie prions. Intraocular inoculation with prions did not provoke scrapie in the graft. These results suggest that the absence of PrPC impairs prion spread from extracerebral sites to the CNS, in addition to blocking neural spread (Brandner et al., 1996b). The neurografting experiments in the PrP null mice further revealed that PrPC-expressing tissues are required for transfer of scrapie infectivity from spleen to the brain (Blättler et al., 1997). Neurografts derived from wild-type or tga20 transgenic mice overexpressing PrP (Fischer et al., 1996) were placed into the brains of Prnpo/o mice. When challenged with scrapie by intraperitoneal or intravenous injection, neither clinical disease nor histopathological changes characteristic of spongiform encephalopathy developed during the observation period, up to 401 days postinoculation (Blättler et al., 1997). Adoptive transfer of PrPexpressing bone marrow cells restored prion replication in the spleen, but did not reconstitute neuroinvasion via the intraperitoneal route. B. Conditional Knockout of Prnp Conventional knockout strategy involves homologous recombination in ES cells and subsequent production of knockouts by injection of the

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modified ES cells into blastocysts. This approach results in early gene inactivation during development, leaving the possibility of compensatory mechanisms masking the possible phenotype of the gene inactivation process. Because the prion gene is active during development, it was reasonable to expect that its inactivation would result in gross abnormalities, but that was not the case as discussed previously. The lack of easily detectable overt phenotype may be a consequence of adaptation during neurodevelopment. There is therefore a need for acute or conditional PrP knockout lines in which the PrP gene can be completely inactivated in adult mice. This need is being addressed in various laboratories; however, a major problem remains—‘leakiness’ of the inducible systems available at present, making it so far difficult to achieve a complete and definitive acute knockout (Tremblay et al., 1998). IV. STRUCTURE AND FUNCTION OF THE PrP GENE After the disruption and successful rescue by complementation of the PrP gene, many research efforts have concentrated on defining ciselements of the PrP gene that are sufficient to support prion pathogenesis and propagation in PrP null mice. Transgenics have facilitated research in this area. A. N-Terminal Deletions Fischer et al. (1996) introduced truncated PrP transgenes encoding PrPC lacking as many as 49 amino-proximal amino acids into PrP knockout mice, and this was sufficient to restore susceptibility to scrapie and support generation of PrPSc in the Zurich I knockout mice. Within the framework of the protein-only hypothesis, these results suggest that the amino-proximal segment of PrPC (including four of the five octarepeats) is not required for conversion into pathogenic infectious form of PrP or for the generation of PrPSc. Under conditions where PrPC is completely digested by proteinase K, PrPSc is partially resistant, having only ~90 N-terminal amino acids cleaved (McKinley et al., 1983) and leaving behind a stable molecule, PrP27–30, capable of supporting prion infectivity. Therefore deletions of up to 49 amino-proximal residues affect only the flexible tail (Riek et al., 1997; Donne et al., 1997) and do not appear to have made enough incursion into the structured globular part, which extends from position 121 to 230 (Riek et al., 1997) to elicit an effect. It is interesting that the octarepeat region of the PrP gene appears from the studies of Fis-

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cher et al. (1996) to be dispensable, as this region has insertional mutations that cause inherited human prion diseases (Collinge et al., 1997) and has also been identified as a copper-binding site of PrP (Hornshaw et al., 1995; Brown et al., 1997a). This initial work was further extended by expressing, in PrP null mice, truncated PrP transgenes with larger deletions extending toward the carboxyl terminus (Shmerling et al., 1998). To ensure correct processing and cellular localization of the truncated PrP genes, the signal peptide and nine additional amino acids were retained in the construct. PrP transgenes lacking residues 32–121 or 32–134 caused severe ataxia and neuronal death limited to the granular layer of the cerebellum as early as 1 to 3 months after birth. However, shorter deletions of 32–80, 32–93, or 32–106 did not show a phenotype (Fischer et al., 1996). This ataxic defect observed in three separate transgenic lines was completely abolished by introducing one or more copies of a wild-type murine PrP gene into mice carrying multiple copies of the truncated PrP gene. The possibility that expression of wild-type PrP could in some way prevent expression of the truncated molecules was ruled out because Western blots of brain extracts showed the same level of truncated PrP in the rescued as in the truncated PrP-expressing mice. Shmerling et al. (1998) explained their observation by speculating that the truncated PrP might be nonfunctional and compete with some other molecule with a PrP-like function for a common ligand. They proposed a model in which truncated PrP acts as dominant negative inhibitor of a functional homolog of PrP, with both competing for the same putative PrP ligand (Shmerling et al., 1998). Incidentally, transmission of brain extract from ataxic truncated PrP-expressing mice to tga20 transgenic mice, that overexpress mouse PrPC did not result in disease for more than 60 weeks after inoculation (Shmerling et al., 1998). When comparing these studies with others, it is worth mentioning that the truncated transgenes of both Fischer et al., (1996) and Shmerling et al., (1998) were based on a mini-Prnp gene, which has been demonstrated to lack expression in Purkinje cells. B. Internal Deletions of PrP Gene Before the availability of Nuclear Magnetic Resonance (NMR) data revealing three α helices in a recombinant PrP (Riek et al., 1997), Muramoto et al. (1997) used a four-helix-bundle model of PrPC (Huang et al., 1994) to construct proteins with each of the predicted helical regions deleted. The aim was to assess the role of each region of the putative secondary structure, as well as those adjacent and intervening

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segments. Fischer et al., (1996) showed in transgenic mice that N-terminal residues 23–88 could be deleted without preventing prion propagation. Using the Mo/SHa chimeric vector MHM2 lacking residues 23–88, already shown by Fischer et al., (1996) to support prion generation, Muramoto et al. (1997) expressed a series of recombinant PrP constructs lacking specific domains in transgenic mice. They observed that although deletion of H1, the STE domain, or the 36-residue segment between H2 and H3 did not produce disease, deletion of H3 or H4 produced a heritable PrP disorder that resembled a neuronal storage disease. There was accumulation of PrP in numerous neurons that were insoluble in nondenaturing detergents, but unlike PrPSc were readily digestible with proteinase-K. This defect was observed in other independent mouse lines with deletions of H3 and H4, and none of the controls with intact H3 and H4 α helices developed CNS dysfunction. Deletion of helices H3 and H4 eliminates a critical region of the hydrophobic core of the prion protein, as well as the disulfide bond (Riek et al., 1996), thus destabilizing the protein (Muramoto et al., 1997). One deletion construct MHM2PrP(∆23–88, ∆141–176) in which residues 23–88 and 141–176 were deleted yielded a PrP fragment of 106 residues that produced protease-resistant PrP in scrapie-infected neuroblastoma (ScN2a) cells (Muramoto et al., 1996). This work was expanded by creating transgenic lines expressing PrP106 on mouse PrP null background (Muramoto et al., 1997). These Tg(PrP106)Prnp o/o mice remained healthy for more than 500 days of age without spontaneous disease. When these transgenic mice, Tg(PrP106)4290/Prnpo/o, were inoculated with full-length, wild-type RML prions that were previously passaged in CD-1 mice, they exhibited neuropathological changes characteristic of experimental scrapie with a mean incubation period of about 300 days (Supattapone et al., 1999). Western blotting demonstrated that the sick Tg(PrP106)Prnp o/o produced proteinase-K resistant PrP106, which migrated as a single diffuse band with an apparent molecular weight of about 22 kDa. Passaging of brain extract from a sick Tg(PrP106)Prnp o/o composed of miniprions containing homologous PrPSc 106 into more Tg(PrP106)Prnp o/o resulted in all inoculated mice developing neurological disease, with a mean incubation period of approximately 66 days, and this reduced incubation period did not change on second passage. The artificial transmission barrier for RML prions containing wildtype PrPSc in Tg(PrP106)4290 mice was dramatically reduced from about 300 days to about 150 days when these mice carried a single copy of wild-type MoPrP gene. The sick Tg(PrP106)4290/Prnp+/o produced both MoPrPSc and PrPSc106 as determined by ELISA using antibodies that distinguish MoPrP-A from PrP106 (Supattapone et al., 1999).

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So why do some deletions of the Prnp gene cause severe ataxia and neuronal loss, whereas others do not? Could a similar mechanism that is proposed to give rise to neuropathology in the ataxic knockout mice also be operating in the transgenics carrying truncated PrP genes? The Prnd gene has been proposed to cause neurodegeneration when upregulated (Moore et al., 1999). Therefore if Prnp and Prnd compete for a common ligand or receptor (Shmerling et al., 1998), then deletions of Prnp that remove the ligand binding domain will remove the inhibitory effect of Prnp on Prnd whose upregulation causes the observed ataxia. C. Genetic Link of PrP Gene to Disease A number of pathogenic mutations in the PrP gene have been linked with disease (see Fig. 1) (for a review see Collinge, 1997). The first mutation to be identified in PRNP was in a family with CJD and constituted a 144 bp insertion into the coding sequence (Owen et al., 1989). A second mutation, a proline to leucine substitution at codon 102 of the human PrP gene, was reported in two families with GSS, and genetic linkage was confirmed between this misense variant and GSS, confirming that GSS was an autosomal dominant mendelian disorder (Hsiao et al., 1989). Inherited prion diseases may produce disease by destabilizing PrPC, which would predispose the molecule to aggregate. Alternatively a mutation could facilitate the interaction between PrPC and PrPSc or affect the binding of a ligand or coprotein. To relate the folding stability of PrPC to its propensity for forming PrPSc, several of the human mutations have been copied into the recombinant mouse protein (Liemann and Glockshuber, 1999). Although this work broadly concluded that there is no absolute correlation between stability and disease, all of the fully penetrant mutations show significant destabilization, whereas polymorphisms have little effect. Other mutations in PRNP have been found to segregate with disorders in families with other varieties of prion diseases, providing evidence for a strong correlation of specific mutations in PrP with different clinical forms of familial prion diseases (Collinge et al., 1989; Medori et al., 1992a; Goldfarb et al., 1991). Pathogenic mutations reported to date in the human PrP gene consist of two groups: (1) point mutations within the coding sequence resulting in amino acid substitutions in PrP or in one case production of a stop codon resulting in expression of a truncated PrP and (2) insertions encoding additional integral copies of an octapeptide repeat present in a tandem array of five copies in the normal protein.

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FIG. 1. Pathogenic mutations identified to date in the human prion gene.

V. TRANSGENIC STUDIES OF PRP TOPOLOGY At the endoplasmic reticulum membrane, the prion protein can be synthesized in several topological forms. The role of these different forms was explored with transgenic mice expressing mutations that alter the relative ratios of the topological forms by Lingappa and coworkers. Previous analysis of PrP topology had suggested that two distinct forms of PrP can be made at the endoplasmic reticulum (ER), one that is fully translocated (secPrP) and one that is transmembrane (Hay et al., 1987a, 1987b; Yost et al., 1990). Digestion of the transmembrane form with proteases added to the outside of the membrane yielded two fragments. One fragment is COOH-terminal derived and glycosylated, and the other is NH2-terminal derived and unglycosylated. Data from Hedge et al., (1998) suggest that NH2- and COOH-terminal fragments reflect the existence of two different transmembrane

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forms of PrP. The C-trans transmembrane (CtmPrP) has the COOHterminus in the ER lumen, with the NH2-terminus accessible to proteases in the cytosol. The other form termed N-trans transmembrane (NtmPrP) has the NH2-terminus in the ER lumen, with the COOH-terminus accessible to proteases in the cytosol. Both transmembrane forms appear to span the membrane at the same hydrophobic stretch in PrP (roughly residues 110 to 135). They both share the epitope for the 3F4 monoclonal antibody (mAb) (to residues 109–112), whereas only the COOH-terminal fragment contains the epitope to the 13A5 mAb (to residues 138–141). These differences in antibody reactivity, glycosylation, and size allow the NtmPrP and CtmPrP fragments to be clearly distinguished (Hedge et al., 1998). Four mutations that altered the ratio of the topological forms when assayed by cell-free translation were identified in preliminary in vitro studies. Two of these mutations (KH→II and ∆STE) were engineered into Syrian hamster PrP (SHaPrP), and the other two (AV3 and G123P) were put into MH2PrP, a mouse-hamster chimera in which residues 94–188 derive from hamster PrP (Scott et al., 1993). The authors observed that the species variation between SHaPrP and MH2PrP (differing at 8 residues) had little effect on topology. However, SHaPrP(KH→II) showed a marked increase (~10 to 50%) in the relative amount of CtmPrP synthesized, with a concomitant decrease in secPrP compared with wildtype SHaPrP (Hedge et al., 1998). The amount of NtmPrP remained unchanged. Similar results were obtained with MH2PrP(AV3) compared with control MH2PrP. In contrast, both SHaPrP(∆STE) and MH2MPrP(G123P) were synthesized exclusively in the secPrP form. Against this background, Hedge et al., (1998) expressed PrP transgenes encoding each of these mutations in mice that lacked the PrP gene FVB/PrP o/o (Prusiner et al., 1993) in order to examine the possible role of PrP topology in neurodegeneration in vivo. These transgenic mice were then observed for clinical signs and symptoms and examined for histopathology. The PrP molecules in their brains were analyzed biochemically for transmembrane topology. Transgenic mice carrying the SHaPrP-(KH→II) mutation developed clinical signs of prion disease, including ataxia and paresis, with an average illness onset of 58 ± 11 days (Hedge et al., 1998). Similarly, mice carrying the MH2MPrP-(AV3) mutation became ill and revealed neurodegeneration with marked astrocytic gliosis (Scott et al., 1997a). However, transgenic mice with high expression levels of SHaPrP(∆STE) and MH2MPrP(G123P) did not show any signs of illness or histological changes, even at ages well beyond the life span of Tg[SHaPrP(KH→II)H], which expressed high levels of the mutant protein.

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The brains of the sick mice were found to contain CtmPrP and not PrPSc. Thus specific transmembrane form of PrP (CtmPrP) can confer severe neurodegeneration in mice with features typical of prion disease (Hedge et al., 1998). The subsequent analysis of the authors of the human neurodegenerative disease caused by A117V mutation (alanine to valine substitution at position 117) revealed that the brain tissue contained CtmPrP, not PrPSc (Hedge et al., 1998). Thus an aberrant regulation of protein biogenesis and topology at the endoplasmic reticulum can result in neurodegeneration. VI. SPONTANEOUS DISEASE IN MUTANT TRANSGENIC MICE Transgenic mice were created to test the biological effects of codon 102 by microinjecting mouse PrP transgene containing a codon 101 leucine substitution (homologous to codon 102 in PRNP). A total of 35 Tg(GSSPrP)174 mice expressing high levels of the mutant transgene product spontaneously developed neurological dysfunction at 166 days of age. (Hsiao et al., 1990). PrPSc levels were low or undetectable, and brain extracts from spontaneously sick mice did not transmit CNS degeneration to wild-type mice. However, transmission to hamsters and Tg(GSSPrP)196 mice expressing lower levels of the homologous mutant transgene product was reported (Hsiao et al., 1994). Given that human 102L cases do transmit to wild-type mice (Tateishi et al., 1996), it is surprising that mouse PrP101L from spontaneously sick Tg(GSSPrP)174 mice failed to transmit to wild-type mice but rather did transmit to hamsters where there is a natural mouse to hamster species barrier to overcome. Another group reported similar spontaneous disease in transgenic mice expressing the mouse homolog of the nine-octapeptide repeat insertion associated with prion disease (Chiesa et al., 1998). The nine octapeptide repeat insertion is the largest known insertion in the PrP gene that is characterized by progressive dementia and ataxia and amyloid plaques in the cerebellum and basal ganglia (Owen et al., 1992; Duchen et al., 1993). These mutant transgenics developed a neurological illness that featured ataxia, neuropathological abnormalities, and the accumulation of PrPSc-like deposits in the brain. The authors observed that elimination of expression of the endogenous wild-type PrP by mating with PrP null mice hastened the onset of symptoms and shortened the clinical phase. A mild degree of protease resistance was reported for the sick mouse brain homogenates, but no proteinase Kresistant material survived the usual digestion conditions (Chiesa et al.,

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1998). Moreover, evidence that these diseases are transmissible was not provided. Several other PrP transgenic mice have been described that spontaneously develop a neurological illness. For instance neurological illness resulting from overexpression of wild-type PrP (Westaway et al., 1994) and deletion transgene constructs (Muramoto et al. 1997; Shmerling et al., 1998) has been described and discussed elsewhere in this chapter. The two examples of P101L and nine octapeptide repeat transgenics did express mutant PrP associated with human inherited prion diseases, although it is difficult to delineate the effects of the respective mutations from simple overexpression effects (Westaway et al., 1994). An alternative approach would be to create point mutations in mice through homologous recombination in ES cells so that only endogenous levels of the mutant protein will be made. Manson et al. (1999) have adopted this approach in creating transgenic mice with P101L (equivalent to P102L in human) mutation in their endogenous PrP. Uninoculated mice both homozygous and heterozygous for the P101L mutation showed no clinical signs of spontaneous TSE for up to 899 days of age. However, these mutant mice showed altered incubation times to disease in response to scrapie infection from different TSE sources when compared to wild-type mice. Manson and co-workers concluded that the 101L PrP gene mutation does not produce any spontaneous genetic disease in mice but significantly alters the incubation time of TSE infection. It is conceivable that if these 101L transgenic mice (Manson et al., 1999) lived long enough to compensate for the low endogenous level of the mutant protein, they might develop spontaneous disease. Such a view is supported by the fact that humans with known pathogenic mutations do not develop disease until late in life. By the same token, overexpression of mutant PrP proteins may accelerate the process of spontaneous conversion, which will manifests as disease in the lifetime of the mice. A number of points about the two P101L experiments described previously are worth noting. First, because the Tg[GSSPrP]196 produced about two times the endogenous level of PrPC (Hsiao et al., 1994) and did not develop spontaneous disease, it is not surprising that the P101L transgenic mice (Manson et al., 1999) expressing only the endogenous level of the mutant protein did not develop spontaneous disease. One potential discrepancy requires further consideration. Although brain homogenate of P101L mice (Manson et al., 1999) that have developed disease following primary inoculation with GSS cases could be passaged to wild-type mice, the Tg[GSSPrP]174 mice (Hsiao et al., 1994), which

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apparently developed a spontaneous prion disease, did not transmit to wild-type mice. This may suggest the generation of different strains of prions in the mice infected with GSS and those developing a spontaneous spongiform encephalopathy. Finally, the fact that the P101L mice (Manson et al., 1999) were more susceptible to human 102L cases than wild-type mice may suggest that the point mutation provided the requisite degree of homology between the host and human inoculum for efficient conversion (Prusiner et al., 1990). VII. TRANSGENIC STUDIES OF THE SPECIES BARRIER A. The Concept of Species Barrier Early experimental transmission studies of human prion diseases were mainly done in laboratory primates, in particular chimpanzees and squirrel monkeys (Brown et al., 1994). These early transmission studies had rather long incubation periods, were expensive, and attracted ethical concerns. Primary transmission of human prions to wild-type laboratory mice have also been fairly unsuccessful, with only occasional transmissions occurring and then at prolonged incubation periods, close to the natural life span of the mice. This difficulty in primary transmissions is the concept of “species barrier” (Pattison and Jones, 1968), the principal determinants of which appear to be degree of homology between the host and the inoculum (Prusiner et al., 1990) and the strain of disease agent. Transgenic mice have contributed enormously in the abrogation of the species barrier, which leads to shortening of incubation time in transmission experiments. The intrinsic properties of the species barrier might result from differences in the primary sequence of PrP of different species. This hypothesis stems from the discovery of the PrP and the subsequent isolation and sequencing of the gene encoding PrP from several species of mammals (Schätzl et al., 1995), including Syrian, Armenia, and Chinese hamsters (Lowenstein et al., 1990). B. Murine to Hamster Species Barrier By splicing together different parts of the coding region or designing transgene constructs with specific mutations, transgenic models with differing properties have been made and are now being used to answer specific questions about prion biology. In a bid to understand the mechanism of species barrier, Scott et al. (1989) made wild-type mice

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transgenic for hamster Prnp and then challenged them with mouse and hamster prions. These transgenic mice, which made endogenous levels of mouse PrPC but differing levels of hamster PrPC, were used to test the hypothesis that the species barrier for transmission of disease is due to differences in the primary structure of PrP. These mice became susceptible to hamster prions and developed disease with a range of scrapie incubation times inversely proportional to the level of expression of hamster PrPC (Scott et al., 1989). Thus the barrier to transmission was abrogated by expression of PrP of the same sequence as that of the inoculating prions, and the efficacy of disease progression was facilitated by increasing the expression level of the homologous PrPC (Prusiner, 1997). On inoculation of the Tg(SHaPrP) transgenics with mouse prions, a prolongation of incubation periods was observed compared with wild-type mice, suggesting that expression of the SHaPrP transgene impeded mouse prion synthesis (Prusiner, 1997). To determine the species range of the prions produced in the transgenic mice, bioassay was used in which brain material from the inoculated animal was further passaged into normal mice or hamsters. Interestingly, the sick transgenic mice would only make prions with the properties of the species in the inoculum (Prusiner et al., 1990). When the mice were inoculated with SHa-adapted Sc237 prions, only SHaPrPSc having characteristics of Sc237 were formed. In contrast, Moadapted RML prions interacted only with MoPrPC to produce neuropathological conditions characteristic of RML scrapie. The implication of these observations is that hamster prions, when presented with both mouse and hamster PrPC, will selectively convert hamster PrPC to hamster PrPSc. Conversely mouse prion will also selectively convert mouse PrPC in these transgenic lines, suggesting that the specificity of prion-host interaction is determined by the primary structure of PrPSc in the inoculum, as well as that of PrPC expressed in the host animal (Prusiner et al., 1990). Therefore the type of PrPSc made in these transgenic mice was dictated by the type of inoculum, in line with the proposal that efficient interaction of PrPSc with host-derived PrPC, required homology between the two interacting molecules (Prusiner et al., 1990). The inverse relationship between gene expression level and incubation period also shows that PrP gene dosage determines the length of incubation time when there is no species barrier to overcome. C. Artificial Prions The preceding example based on wild-type prion protein is corroborated by results from transgenic mice producing artificial PrPC. Scott et

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al., (1993) created transgenic lines in which the central portion of the mouse gene was replaced with the equivalent region of hamster prion protein gene. Two SHa/MoPrP transgenic lines were produced in which the first, Tg(MH2M), contains five amino acid substitutions encoded by SHaPrP; and the other, Tg(MHM2), has two substitutions. The normal SHaPrP and MoPrP genes differ at 16 positions within the open reading frame (ORF). Mice from these two transgenic lines producing chimeric PrPC were then challenged with Syrian hamster (Sc237) or mouse scrapie (RML) prions. Both lines were susceptible to infection with Mo(RML) prions, but only MH2M was susceptible to SHa(SC237) prions (Scott et al., 1993). The two amino acid substitutions in the MHM2 were not enough to abrogate the species barrier between mice and hamsters. When the resulting prions designated MH2M(Sc237) were passaged in mice and hamsters, they were able to infect both Syrian hamsters and mice. More important they showed a distinct preference for infection of mice expressing the homologous MH2M PrPC. This preference for infection of animals expressing a homologous PrPC supports the view that there is a recognition and homophilic interaction of PrPSc molecules in the inoculum with the PrPC in cells of the host. Of interest, whereas Mo(RML) prions do not ordinarily infect Syrian hamsters, inoculation of brain homogenates from Tg(MH2M) mice into Syrian hamsters had a 100% attack rate, indicating that MH2M(RML) had been formed. The sick hamsters had a unique pattern of PrPSc accumulation, suggesting that the artificial prions created by the chimeric transgene were a new prion isolate. D. Human to Murine Species Barrier In general, primary transmission from one species to another occurs much less efficiently, if at all, than transmission within the same species. Early attempts to transmit human prions to transgenic mice met with varied success. The Tg(HuPrP)110 and Tg(HuPrP)152 transgenic lines were made by inserting wild-type human PrP coding sequences encoding valine at codon 129 into cosSHaTet vector (Telling et al., 1994). When these transgenic mice were inoculated with brain homogenates from cases of GSS and sporadic and iatrogenic CJD cases, only 14 of 169 exhibited scrapie-like symptoms with protracted incubation times of > 589 days. A similar percentage of nontransgenic controls became sick with similar incubation times. The authors concluded that additional species-specific factors might be involved in prion replication. However, exceptions to this finding were reported by Collinge et al. (1995a).

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These authors inoculated a panel of human CJD cases into the Tg(HuPrP)152 transgenic mice and showed that one human CJD case, designated PDG170, resulted in eight of nine mice dying from neurological disease, with a mean incubation time of 297 days. Following the apparent lack of susceptibility of the Tg(HuPrP), Telling et al. (1994) then constructed mice that expressed a chimeric prion protein in which a segment of mouse PrP was replaced with the corresponding human PrP sequence. Whereas HuPrP differs from MoPrP at 28 of 254 residues, the chimeric PrP designated MHu2MPrP differed from MoPrP by only nine amino acids between residues 96 and 167. When similarly inoculated with CJD prions, all the Tg(MHu2MPrP) mice encoding methionine at codon 129 developed neurological disease ~200 days after inoculation with brain homogenates from three CJD patients, all homozygous for methionine at polymorphic residue 129. While the chimeric transgene approach (Telling et al., 1994) was being pursued, the endogenous mouse PrP allele was also removed by breeding the Tg(HuPrP)152 mice to PrP null mice (Prnp o/o) to produce homozygous Tg(HuPrP)152/Prnp o/o. It is clear that breeding out the endogenous mouse PrP alleles has made the Tg(HuPrP)152/Prnp o/o mice highly susceptible to CJD, with all inoculated mice succumbing at short incubation periods usually in the range of 180 to 220 days (Collinge et al., 1995a; Telling et al., 1995). In contrast, the Tg(MHu2MPrP) mice were rendered only slightly more susceptible when the endogenous mouse PrP gene was removed (Telling et al., 1995). The Tg(HuPrP)152/Prnp o/o mice (homozygous for the human transgene and without endogenous mouse PrP) are susceptible to all isolates of classic human CJD, and there is no decrease in incubation period on primary and second passage (Collinge et al., 1995a). All of 16 CJD cases encompassing a wide range of clinicopathological phenotypes, all three PrPSc types reported in sporadic and iatrogenic prion diseases (Collinge et al., 1996), and all PRNP genotypes at polymorphic codon 129 have been transmitted to the Tg(HuPrP)152/Prnp o/o transgenic mice (Hill et al., 1997). Important data on prion strains have been generated using these transgenic mice and are discussed later in this chapter. The different susceptibilities observed between Tg(HuPrP)152 and Tg(MHu2M) mice led to the proposal of the existence of an additional protein, provisionally designated protein X, that might be necessary for susceptibility of the Tg(HuPrP) mice to human prions (Telling et al., 1995). The authors suggested that protein X might function as a molecular chaperone required in the formation of nascent PrPSc. Telling et al. (1995) suggested that MoPrPC produced by Tg(HuPrP)152 inhibited the conversion of HuPrPC into PrPSc and that inhibition was abol-

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ished once MoPrPC was removed by gene ablation. They reasoned that the MoPrPC must be binding to mouse protein X with a considerably higher affinity than does HuPrPC, thus providing an explanation for why MoPrPC inhibits the transmission of human prions in Tg(HuPrP)152 (Telling et al., 1995). Because the Tg(MHu2M) were found to be susceptible to human prions irrespective of the presence of endogenous MoPrPC, the binding site for protein X was proposed to be in the C-terminal end of PrPC within the mouse encoded residues of the chimeric PrPC transgene construct, which was missing from the Tg(HuPrP)152 construct (Telling et al., 1995). The authors proposed that the mouse sequences in the chimeric PrPC enabled it to compete effectively with MoPrPC for binding to protein X. Although it is to be anticipated that other cellular cofactors will affect the efficiency of prion propagation, there remains no direct evidence for the existence of protein X. It is possible that the transgenetic observations that led to this hypothesis could be explained by mouse PrP itself acting to interfere with interactions between human PrPSc and human PrPC. Of interest, mice expressing a chimeric mouse/bovine PrP, which would be expected according to the protein X hypothesis to be more sensitive to bovine prions than mice expressing wild-type bovine PrP in the presence of mouse PrP, are in fact much less susceptible (see later). E. Bovine to Murine Species Barrier Understanding what constitutes the species barrier has important implications for assessing the risk of BSE transmission to humans. Following the success in transmitting human prions to the chimeric Tg(MHu2M) transgenics (Telling et al., 1994), transgenic mice were created that expressed either an intact bovine ORF (BoPrP) or a bovine/mouse chimeric ORF (MBo2M) in the cosTet vector (Scott et al., 1997b). On inoculation of Tg(MBo2M)Prnp o/o and Tg(BoPrP)Prnp o/o mice with bovine prions, none of the former group of mice developed neurological dysfunction more than 600 days after inoculation. This was contrary to the authors’ expectation, as the MBo2M ORF was composed of BoPrP and MoPrP-A sequences, and their previous studies had shown that Prnpa mice were susceptible to BSE. The same inoculum was able to transmit disease to control mice including FVB, Tg(BoPrP)833, and Tg(BoPrP)333 mice expressing MoPrP-A. The Tg(BoPrP)833 and Tg(BoPrP)333 were found not to express bovine PrP at levels detectable by Western blotting and were therefore used as Prnpa/a controls. When Tg(BoPrP)4125/Prnp o/o mice expressing high levels of BoPrP were

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inoculated with BSE, they exhibited clinical signs within 250 days. A second line Tg(BoPrP)4092/Prnp o/o with an intermediate level of BoPrP expression had a longer incubation period with a mean of about 320 days (Scott et al. 1997b). Primary passage of three separate cases of vCJD killed Tg(BoPrP) 4125/Prnp o/o mice with incubation periods of 250 to 270 days, comparable to that obtained with cattle BSE and secondary passage of BSE from Tg(BoPrP)4125/Prnp o/o mice (Scott et al., 1999). This is perhaps not surprising, as vCJD has been shown to share both the molecular and biological properties of the BSE strain. These data are in accord with the clear evidence that vCJD is caused by the same agent strain as BSE (Collinge et al., 1996; Bruce et al., 1997; Hill et al., 1997). However, the bovine to human species barrier, which is discussed further later, was not directly tested by Scott et al. (1999). The Tg(BoPrP)4125/Prnp o/o mice were also susceptible to natural sheep scrapie deriving from Suffolk sheep in which susceptibility to scrapie is controlled by homozygosity for glutamine (Q) at the PrP codon 171 polymorphism (Hunter et al., 1997). Bovine PrP contains Q at the equivalent position in sheep PrP; thus the ease of transmission of scrapie to Tg(BoPrP) may be due to the homology at this polymorphic residue implicated in susceptibility of sheep to scrapie. To develop an improved bioassay for the detection of cattle-derived BSE prions, Buschmann et al. (1999) generated transgenic mice overexpressing bovine (Tgbov XV) or murine/bovine chimeric PrPC (Tgmubo XIII). They then studied their susceptibility together with conventional RIII and murine PrPC overexpressing tga20 mice, by inoculating the four groups of mice with cattle-derived BSE prions. Although overexpression of murine PrPC in tga20 mice did not lead to a shorter incubation time than conventional RIII mice, the Tgbov XV mice showed markedly reduced incubation times, with a mean of about 250 days. Like the chimeric Tg(Bo2M)Prnp o/o of Scott et al. (1997b), the Tgmubo XIII chimeric mice displayed an extremely low susceptibility to BSE as no animal showed clear clinical symptoms even after 800 days. Brains of sick Tgbov XV mice had histopathology and glycoform band patterns typical of BSE. Secondary passage of brain homogenate of sick Tgbov XV did not lead to a lowering of incubation period, indicating that expression of the bovine sequences alone were sufficient to abrogate the species barrier between bovine and mice. Scott et al. (1997b) do offer a possible explanation for the different susceptibilities of the two chimeric transgenic mice, Tg(MHu2M) and Tg(MBo2M). A comparison of the MoPrP-A with MBo2M PrP and MHu2M PrP translated sequences shows that human residue substitu-

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tions in MHu2M extend from 97 to 168, whereas Bo substitutions in MBo2M extend from 97 to 186. This finding raised the possibility that residues 184 and 186, which are not homologous in Bo and Mo PrP and lie at the C-terminal end of the chimeric region, might account for the differences in susceptibility of Tg(MHu2M) and Tg(MBo2M) mice to prion infection (Scott et al., 1997b). Alternatively, residue 203, which is a Val in Mo and Hu PrP and is Ile in BoPrP, might be responsible for this difference in susceptibility to prions. In Tg(MBo2M) mice, residue 203 is a Val and thus might prevent conversion of MBo2M PrPC into PrPSc (Scott et al., 1997b). F. Bovine to Human Species Barrier The emergence of a new variant form of CJD in young people (vCJD) in the United Kingdom raised the possibility that BSE might have crossed the species barrier between humans and cattle. There is compelling evidence that vCJD and BSE are the same strain (Collinge et al., 1996; Bruce et al., 1997; Hill et al., 1997). What is not so clear is the magnitude of the bovine to human species barrier. Clearly, this cannot be measured directly, as this would require inoculation of humans with BSE. However, transgenic models may offer a way to address this issue, at least in part. Early studies of the molecular basis of the species barrier suggested that it mainly resided in differences in PrP primary structure between the species from which the inoculum was derived and the inoculated host. The fact that most sporadic and acquired CJD occurred in individuals homozygous at PRNP polymorphic codon 129, supported the view that prion propagation proceeded most efficiently when the interacting PrPSc and PrPC were of identical primary structure (Collinge et al., 1991). However, it has long been recognized that the type of prion strain affects ease of transmission to another species. With BSE prions, the strain component of the barrier seems to predominate, with BSE not only transmitting efficiently to a range of species, but maintaining its transmission characteristics even when passaged through an intermediate species with a distinct PrP gene (Bruce et al., 1994). For example transmission of CJD prions to conventional mice is difficult, with few if any inoculated mice succumbing after long incubation periods, which is consistent with a substantial species barrier (Hill et al., 1997; Collinge et al., 1995a). In sharp contrast, transgenic mice expressing only human PrP Tg(HuPrP)152/Prnp o/o are highly susceptible to CJD prions, with 100% attack rate and consistent short incubation periods that are unaltered by second passage, consistent with a complete lack of species barrier

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(Collinge et al., 1995a). However, variant CJD prions (human PrP of identical structure) transmit much more readily to wild-type mice than do classic CJD prions, whereas transmission to transgenic mice is less efficient than with classic CJD (Hill et al., 1997). The term “species barrier” does not seem appropriate to describe such effects, and species-strain barrier or transmission barrier may be preferable (Collinge, 1999). PrP amino acid sequence and strain type affect the three-dimensional structure of glycosylated PrP that will presumably, in turn, affect the efficiency of the protein-protein interactions thought to determine prion propagation. Contribution of other components to the species barrier is possible and may involve interaction of cofactors that mediate the efficiency of prion propagation, although no such factors have yet been identified. The species barrier between cattle BSE and human beings can be modeled in transgenic mice expressing human PrPC, which produce human PrPSc when challenged with human prions (Collinge et al., 1995a). When such mice—expressing human PrP valine 129 (at high concentrations) and mouse PrP—are challenged with BSE, three possibilities may occur: The mice could produce human prions, murine prions, or both. In fact, only mouse-prion replication could be detected. Although there are caveats for this model, especially that propagation of human prions in mouse cells may be less efficient than that of mouse prions, this result would be consistent with the bovine-human barrier being larger than the bovine-mouse barrier for this PRNP genotype. In the second phase of these experiments, mice expressing only human PrP were challenged with BSE. Although CJD isolates transmit efficiently to such mice at around 200 days, only infrequent transmissions at more than 500 days were seen with BSE, consistent with a substantial species barrier for this human PRNP genotype. The PRNP valine 129 genotype was studied initially in attempts to produce an animal model of human prion disease, as this genotype was overrepresented among early cases of iatrogenic CJD (Collinge et al., 1991), which suggests increased susceptibility or shorter incubation periods in this genotype. These studies are being repeated in mice expressing only human PrP methionine 129 and in heterozygotes. So far, vCJD has affected approximately 75 people in the United Kingdom and 2 in France, and to date all these cases have the PRNP codon 129 methionine homozygous genotype. VIII. TRANSGENIC STUDIES OF INCUBATION PERIOD Incubation period is the time between experimental inoculation of scrapie prions to diagnosis of neurological dysfunction based on prede-

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termined neurological signs. This property has been used to characterise prion strains and was initially thought to be controlled by a distinct locus variously called Sinc or Prn-i. The two PrP alleles that are found in short and long incubation time mice based on ME7 inoculation are designated Prnpa and Prnpb, respectively. These two alleles differ by two amino acid residues at dimorphic positions 108 (leucine-phenylalanine) and 189 (threonine-valine) (Westaway et al., 1987), with both of the PrPB mutations (108F 189V) found only in inbred mouse strains that have long incubation times (Carlson et al., 1988, Westaway et al., 1987). Early attempts by backcross of NZW x I/LnJ to confirm that Prn-i might be congruent with Prn-p were not successful. Individual mice with discordant scrapie incubation time and Prn-p genotype were observed, suggesting meiotic recombination between Prn-i and Prn-p (Carlson et al., 1986; Race et al., 1990). In an attempt to clarify the relationship between Prn-i and Prn-p Westaway et al. (1991) resorted to transgenics. The long incubation time allele of Prn-i is dominant and so it was expected that expressing Prn-pb in short incubation mice (Prn-pa) would lead to prolonged incubation times. Contrary to expectation, mice from four independent Tg(Prn-pb) lines showed shorter scrapie incubation times than nontransgenic control animals (Westaway et al., 1991). These results originally described as paradoxical were dictated by gene dosage of the Prn-pb transgene, thus masking genuine congruence of Prn-p/Prn-i. Moore et al. (1998) used gene targeting to try to resolve these difficulties surrounding the identity of Sinc/Prn-i and Prn-p. They constructed co-isogenic 129/Ola mice, which differed only by targeted codon 108 and 189 using a twostep double replacement gene targeting strategy. The 129/Ola mice expressing the gene-targeted PrP-B allotype instead of PrP-A had dramatically shortened incubation times when infected with the mouseadapted BSE strain 301V, similar to the short incubation times of VM/Dk (Sincp7/p7) mice (Moore et al., 1998). Whereas mice homozygous for the targeted allele (Prnp-A, 108F 189V) had an average incubation time of 133 days, the wild-type 129/Ola mice (Prnp-A) had incubation time of 244 days. Thus data from gene-targeting studies provided evidence that the PrP gene itself is responsible for differences in incubation periods. It is perhaps worth noting that the original characterisation of the short (Prnpa) and long (Prnpb) incubation time alleles have been based on the ME7 strain (Dickenson et al., 1968). Moore et al. (1998), however, used 301V, which is a mouse passaged BSE in their study. That only two amino acid differences at codon 108 and 189 can cause such a dramatic alteration in incubation periods demonstrates that Sinc/Prn-i and the PrP gene are congruent. It remains to be seen,

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however, whether ME7 inoculation into these mice would lead to the same or a different set of results to that observed for 301V. IX. TRANSGENIC STUDIES OF THE MOLECULAR BASIS OF PRION STRAINS A. What Are Strains? Distinct prion isolates are referred to as strains. Strains of agent are defined and identified by their biological properties such as pattern of the incubation periods, lesion profile (the distribution of pathological change in the brain) in mice of defined PrP genotype (Bruce et al., 1991), and by behavioural and metabolic differences in the host (Carp et al., 1994). The existence of multiple strains has been difficult to accommodate within the protein-only hypothesis and has led to the conclusion that the disease agents carry some form of strain-specific information that determines disease characteristics. In rodents with experimental prion disease, the neuroanatomic pattern of pathology, so-called lesion profile, is thought to be largely a combination of host and strain effects. Biological diversity is generally encoded within nucleic acids; yet no chemical, biological, or physical evidence argues in favor of scrapie-specific polynucleotide (Prusiner, 1991). The gene-targeting studies discussed earlier have clearly established that the endogenous PrP gene may determine how the host responds to a particular scrapie inoculum. However, the ability of different scrapie isolates or strains to maintain their respective properties through repeated passage in the same mouse strain homozygous for the PrP gene cannot be explained by differences in primary structure. Furthermore, strains can be reisolated in mice after passage in intermediate species with different PrP primary structures (Bruce et al., 1994). B. Molecular Strain Types Recently, several human PrPSc types have been identified that are associated with different phenotypes of CJD (Collinge et al., 1996; Parchi et al., 1996). The different fragment sizes seen on Western blots after treatment with proteinase K digestion suggest that there are several different human PrPSc conformations. However, to fulfil the criteria of strains, these patterns must be transmissible to animals both in the same and in different species. Remarkably, this is the case, with both PrPSc fragment sizes and the ratios of the three PrP glycoforms (digly-

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cosylated, monoglycosylated, and unglycosylated PrP) maintained on passage in transgenic mice expressing human PrP (Collinge et al., 1996). Transmission of human prions and bovine prions to wild-type mice results in murine PrPSc with fragment sizes and glycoform ratios that correspond to the original inoculum (Collinge et al., 1996). Using the Tg(HuPrP)152/Prnp o/o mice discussed previously as experimental models, Collinge et al. (1996) have reported that transmission of type-1 CJD, which are all genotype MM, resulted consistently in a type-2 banding pattern of human protease-resistant PrP being produced in the mice. The phenomenon of strain mutation has long been recognized in conventional strain-typing studies. For example, when cloned scrapie strain 22A, which is stable in Sincp7 genotype, is passaged several times in SincS7 genotype, a new strain, 22F, with altered incubation period characteristics is generated (Bruce, 1993). Remarkably, transmission of type-2 CJD (of all three codon-129 genotypes) or type-3 CJD (which were all genotype MV or VV) resulted in production of protease-resistant human PrP in the Tg(HuPrP)152/Prnp o/o mice with an identical banding pattern to the primary inoculum (type 2 or 3, respectively). However, a study of mean incubation periods between inocula of different 129 genotypes did not show any significant difference (Collinge et al., 1996). It has previously been shown that the PrPSc type seen in vCJD (type 4) is distinct from those seen in classic CJD and has a ratio of glycoforms closely similar to that of BSE passaged in several other species (Collinge et al., 1996). Of interest, when vCJD was transmitted to these Tg(HuPrP)152/Prnp o/o mice (carrying valine at polymorphic residue 129), a new strain designated type 5 was produced. The type 5 shares the glycoform ratio of type 4, but the fragment sizes differ from those in the inoculum and were indistinguishable from those in the type-2 pattern (Hill et al., 1997). The production of a distinct molecular strain type on transmission of vCJD to mice expressing valine 129 human PrP suggests that BSE transmitted to humans of this genotype might produce a similar strain. Such cases may differ in their clinical and pathological phenotype to vCJD, but could be identified by PrPSc typing. These data suggest that strain type can be as important as PrP primary structure differences between donor and host in species barrier. The data also strongly support the “protein-only” hypothesis of infectivity and suggest that strain variation is encoded by a combination of PrP conformation and glycosylation. Transmission of PrPSc fragment sizes from two different subtypes of inherited prion disease to transgenic mice expressing a chimeric human mouse PrP has also been reported (Telling et al., 1996). These

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transmission studies in mice carrying valine at codon 129 of the PrP gene resulting in strain mutations (Bruce, 1993) need to be repeated in human PrP transgenic mice carrying methionine at codon 129 of the prion gene. This will clarify the effect of codon 129 mismatch on transmission characteristics of type-1 CJD, BSE, and vCJD. Such experiments are ongoing (Asante et al., unpublished). As PrP glycosylation is thought to be a cotranslational process, the different glycoform ratios may represent selection of particular PrPC glycoforms by PrPSc of different conformations (Collinge et al., 1996). According to such a hypothesis, PrP conformation would be the primary determinant of strain type, with glycosylation being involved as a secondary process. However, because different cell types may glycosylate proteins differently, PrPSc glycosylation patterns may provide a substrate for the neuropathological targeting that distinguishes different prion strains. Particular PrPSc glycoforms may replicate most favorably in neuronal populations with a similar PrP glycoform expressed on the cell surface. Such targeting could also explain the different incubation periods, which also discriminate strains, targeting of more critical brain regions, or regions with higher levels of PrP expression, producing shorter incubation periods. Such studies may allow a new molecular classification of human prion diseases; it is likely that additional PrPSc types or strains will be identified. This may well open new avenues of epidemiological investigation and offer insights into causes of “sporadic” CJD. In addition, PrPSc typing can be applied to other species; it is already apparent that PrP glycoform analysis alone can distinguish a number of mouse-passaged scrapie strains (Somerville et al., 1997). The combination of fragment size and glycoform analysis should allow better resolution and might be applied, for instance, to study if BSE has transmitted to other species involved in the human diet such as sheep (Collinge et al., 1996) The SHaPrP is glycosylated at Asn residues 181 and 197 (Endo et al., 1989), and sugar chains have been shown to modify the conformation and interaction of some glycoproteins (O’Connor and Imperiali, 1996). DeArmond and co-workers postulated that variations in complex CHOs may alter the size of the energy barrier that must be traversed during formation of PrPSc. If this were the case, then regional variations in CHO structure in the CNS could account for selective formation of PrPSc in particular areas of the brain (DeArmond et al., 1997). They tested this postulate by making transgenic mice expressing PrPs mutated at one or both of the glycosylation consensus sites. These mutations resulted in aberrant topologies of PrPC within the CNS. Dele-

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tion of the first oligosaccharide altered PrPC trafficking and prevented infection with two prion strains. Deletion of the second did not alter PrPC trafficking, permitted infection with one prion strain, and had a profound effect on the PrPSc deposition pattern. The procedure for altering the glycosylation sites required the substitution of an Ala for Thr and so the possibility of the introduced amino acid substitution confounding the effects of the loss of CHO was addressed (DeArmond et al., 1997). The authors examined the effects of other mutations known to modify the extent of glycosylation and to cause inherited prion diseases (Monari et al., 1994; Parchi et al., 1995) and found that the point mutations introduced at the consensus sites did not alter the distribution of PrPC. They concluded that glycosylation could modify the conformation of PrPC and affect the affinity of PrPC for a particular conformer of PrPSc, thereby determining the rate of nascent PrPSc formation and the specific patterns of PrPSc deposition. The ability of a protein to encode a disease phenotype has important implications in biology, as it represents a nonmendelian form of transmission. It would be surprising, and also itself intriguing, if evolution had not used this mechanism for other proteins in a range of species. The identification of prion-like mechanisms in yeast is particularly interesting in this regard (Wickner, 1994; Ter Avanesyan et al., 1994). C. What Is the Cause of Neurodegeneration in Prion Diseases? Whether PrPSc causes disease through neurotoxicity by accumulating in neuronal cells, whether the depletion of PrPC causes disease, or whether some other unknown mechanism is responsible is unclear. Data from neurografting experiments (Brandner et al., 1996a, 1996b) led to the implication that it is not deposition of PrPSc but rather the availability of PrPC for some intracellular process elicited by the infectious agent that is directly linked to spongiosis, gliosis, and neuronal cell death (Brandner et al., 1996a, 1996b). In vitro studies have shown that the activity of Cu/Zn superoxide dismutase is reduced in Prnpo/o mice, suggesting that PrPC may be important for cellular resistance to oxidative stress (Brown et al., 1997b). Abnormalities of synaptic inhibition seen in PrPC-deficient mice (Collinge et al., 1994), which mimic neurophysiological abnormalities seen in patients with CJD and in scrapie-infected mice (Jefferys et al., 1994), argue in favor of loss of function. However, the observation that PrPC-overproducing transgenic mice succumb to disease rapidly, but relatively little of it is converted to PrPSc might counteract the loss of function argument (Fischer et al., 1996). Also the fact that Prn-p o/o mice

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are normal in their development and behavior has been used by proponents of toxic gain of function to argue that scrapie pathology is due to intrinsic neurotoxicity of PrPSc. Furthermore, deposition of PrPimmunoreative material colocalizes with typical histopathology, and synthetic amyloidogenic PrP fragments as well as liposome-packaged PrP27–30 (the protease-resistant core of PrPSc) are neurotoxic in vitro (Forloni et al., 1994). The reported neurotoxicity of PrP106–126 in cultured cells has recently been called into question by Kunz et al. (1999). The authors found that PrP106–126 concentrations of up to 200 µM did not show any toxic effect on cultured mouse cortical neurons. They ascribed the reported toxic effect of PrP106–126 to an unidentified contaminant resulting from the synthesis of the peptide or the highperformance liquid chromatography (HPLC) purification rather than the peptide itself (Kunz et al., 1999). The same is true for toxic gain of function. That is, the notion that PrPSc accumulation directly causes disease has not been proven conclusively because of reports that mice succumbing to prion disease may be devoid of detectable PrPSc (Hsiao et al., 1994). PrPSc does not appear to be toxic to cells lacking PrPC (Brandner et al., 1996a), and PrPSc is difficult to detect in some cases of inherited human prion diseases such as fatal familial insomnia (FFI) (Medori et al., 1992b), as well as in transgenic mice infected experimentally with FFI (Collinge et al., 1995b). These examples suggest that, at least in these cases, neurodegeneration may not be the result of PrPSc toxicity. Furthermore, although all C57BL/6 mice inoculated with BSE-infected cattle brain homogenate exhibited symptoms of neurological disease, more than 55% of the sick mice had no detectable PrPSc (Lasmezas et al., 1997). Although neurological examination revealed neuronal death in all mice, classic TSE-associated changes such as neuronal vacuolation and astrocytosis were limited to sick mice with PrPSc deposition. Subsequent passage of brain material from PrPSc-negative mice transmitted disease, and by the third passage almost all sick mice had PrPSc deposition and typical TSE pathology, a consequence of adaptation in the new host. The authors concluded that although PrPSc has a clear role in TSE, it may not be the transmissible component of the infectious agent, and it may be involved in species adaptation. Büeler et al. (1994) observed that transgenic mice having a single copy of the mouse prion gene (Prn-po/+) had incubation times of about 41 weeks, but at about 24 weeks they already had high levels of PrPSc in the brain, but were still months away from showing scrapie symptoms. The development of gliosis, spongiosis, and loss of CNS neurons were only marginally delayed in the Prn-po/+ mice compared with the wild-

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type mice and was largely dissociated from the clinical condition of the mice at later time points. The authors suggested that, based on these finding status spongiosis and astrogliosis might not be the main factor leading to clinical disease in mouse scrapie. If gain of function is the mode of disease development, however, then the findings of Büeler et al. (1994) may suggest a critical threshold of PrPSc accumulation below which mice are tolerant. Alternatively, the gain of function implied by the autosomal dominant mode of inheritance could reflect the formation of PrPSc, which then functionally depletes PrPC by a dominant negative effect (Collinge, 1997). X. TRANSGENE VECTOR CONSIDERATIONS The presence of at least one intron is essential for the correct and efficient expression of transgenes (Brinster et al., 1988), and the prion gene, notwithstanding its unique biological properties, is no exception. The hamster and human PrP genes are made of two exons separated by a 10 kb intron (Basler et al., 1986; Kretzschmar et al., 1986), with exon 2 encoding the ORF and 3′ untranslated region. The mouse, bovine, and sheep PrP genes contain three exons and two introns, with exon 3 being analogous to exon 2 of the hamster PrP gene. The upstream intron of mouse PrP is 2 kb, and the downstream intron is 6 to 12 kb (Westaway et al., 1994). The early PrP transgenes were based on a 40-kb cosmid vector (SHaCosTt) containing genomic hamster Prnp 5′ and 3′ elements of hamster PrP (Scott et al., 1989; Hsiao et al., 1990). Fischer et al. (1996) generated PrP-encoding constructs in which the large intron (half-genomic construct or phgPrP) or both introns (cDNA construct or pPrPcDNA) were deleted. These constructs and the Prnp-containing cosmid DNA, cos6.I/lnJ4 (Westaway et al., 1994), were introduced into Prnp o/o or Prnp o/+ mice by pronuclear injection. In five of six half-genomic transgenic lines and one cosmid transgenic line, there was PrP expression as monitored by Northern analysis; however, of eight lines tested for the cDNA construct, none, including those with >150 copy numbers, showed detectable levels of PrP protein in the brain and very weak to no mRNA expression. (Fischer et al., 1996). An interesting result of removing the large intron 2 of MoPrP was that PrP RNA was not detectable in Purkinje cells of the half-genomic mice, but was abundant in Purkinje cells of wild-type and cosmid transgenic mice. The authors suggested that one or more control elements responsible for the specific expression of PrP in Purkinje cells (presumed to be resident in the large intron) might be absent from the halfgenomic construct.

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Furthermore, whereas animals expressing half-genomic PrP-A at higher levels than the cosmid transgenics revealed no obvious pathological phenotype, some animals from the latter group developed paresis of the their hind-limbs as early as 4 to 6 months. Similar symptoms in Prnp cosmid transgene overexpressing mice have been described (Westaway et al., 1994) and could be due to differences in expression pattern or overexpression of an unidentified gene product encoded by sequences present in the cosmid, but not in the half-genomic construct (Fischer et al., 1996). Although others (Chiesa et al., 1998) using a similar half-genomic vector have also reported absence of expression in Purkinje cells in transgenics, heterologous promoters, and introns have been used to efficiently express hamster PrP in transgenic mice. Race and co-workers created hamster PrP expressing transgenic mice (Tg52NSE) using the rat neuron specific enolase promoter and its intron (Race et al., 1995). Not only was there expression in cerebellar Purkinje cells, the mice that expressed HaPrPC only in neurons were susceptible to intracerebral inoculation of hamster scrapie strain 263K. There was no expression in glial cells or cells within the spleen or lymph nodes, suggesting that expression of PrPC in non-neuronal brain cells, including astrocytes, is not necessary to overcome the TSE species barrier (Race et al., 1995), at least where infection is via the intracerebral route. However, experiments in PrP-null mice that were neurografted and subsequently challenged intraocularly with PrPSc showed that PrPC is necessary for prion spread along the neural pathways (Brandner et al., 1996b). Reports that astrocytes and splenic follicular dendritic cells (FDC) may be the earliest sites of PrPSc accumulation following scrapie infection (Diedrich et al., 1991, Moser et al., 1995) may be more relevant to peripheral inoculations. It would be interesting therefore to examine how the Tg52NSE mice lacking expression in the lymphoreticular system would respond to peripheral inoculation with hamster strain 263K. The data of Race et al. (1995) also suggest that expression of PrP in Purkinje cells is not the preserve of the large intron of the prion protein gene. It had been suggested that Purkinje cell-specific enhancer might be contained within the second large intron (Fischer et al., 1996). Race et al. (1995) concluded that susceptibility of their mice was mediated by the 1 kb HaPrP cDNA, including the ORF rather than the additional 39 kb of transgene DNA used in previous studies (Scott et al., 1989; Westaway et al., 1994). A mini human PrP promoter reporter construct using only a 290 bp generic intron has successfully targeted expression to neuronal cells including Purkinje cells (Asante et al., unpublished).

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XI. CONCLUDING REMARKS Transgenic technology has made an invaluable contribution to acquisition of new knowledge in prion biology, particularly in structural studies. A number of important questions about prion biology cannot be answered in a cell culture system and require the use of an animal model. For instance, the search for the bonafide function of PrP gene is likely to be resolved through transgenic and knockout technology. ACKNOWLEDGMENT We thank Graham S. Jackson and Eckhard Flechsig for helpful discussions and access to information.

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Tobler, I., Gaus, S.E., Deboer, T., Achermann, P.A., and Manson, J.C. (1996). Altered circadian activity rhythms and sleep in mice devoid of prion protein. Nature (Lond.) 380, 639–642. Tremblay, P., Meiner, Z., Galou, M., Heinrich, C., Petromilli, C., Lisse, T., Cayetano, J., Torchia, M., Mobley, W., Bujard, H., DeArmond, S. J., and Prusiner, S.B. (1998). Doxycycline control of prion protein transgene expression modulates prion disease in mice. Proc Natl Acad Sci USA 95, 12580–12585. Westaway, D., Goodman, P.A., Mirenda, C.A., McKindly, M.P., Carlson, G.A., and Prusiner, S.B. (1987). Distinct prion proteins in short and long scrapie incubation period mice. Cell 51, 651–662. Westaway, D., Mirenda, C.A., Foster, D., Zebarjadian, Y., Scott, M., Torchia, M., Yang, S.L., Serban, H., DeArmond, S.J., Ebeling, C., Prusiner, S.B., and Carlson, G.A. (1991). Paradoxical shortening of scrapie incubation times by expression of prion protein transgenes derived from long incubation period mice. Neuron 7, 59–68. Westaway, D., DeArmond, S.J., Cayetano, C.J., Groth, D., Foster, D., Yang, S.L., Carlson, G.A., and Prusiner, S.B. (1994). Degeneration of skeletal muscle, peripheral nerves, and the central nervous system in transgenic mice overexpressing wild-type prion proteins. Cell 76, 117–129. Wickner, R.B. (1994). [URE3] as an altered URE2 protein: evidence for a prion analog in Saccharomyces cerevisiae. Science 264, 566–569. Will, R.G., Ironside, J.W., Zeidler, M., Cousens, S.N., Estibeiro, K., Alperovitch, A., Poser, S., Pocchiari, M., Hofman, A., and Smith, P.G. (1996). A new variant of CreutzfeldtJakob disease in the UK. Lancet 347, 921–925. Whittington, M.A., Sidle, K.C.L., Gowland, I., Meads, J., Hill, A.F., Palmer, M.S., Jefferys, J.G.R., and Collinge, J. (1995). Rescue of neurophysiological phenotype seen in PrP null mice by transgene encoding human prion protein. Nat. Genet. 9, 197–201. Yost, C.S., Lopez, C.D., Prusiner, S.B., Myers, R.M., and Lingappa, V.R. (1990). Nonhydrophobic extracytoplasmic determinant of stop transfer in the prion protein. Nature 343, 669–672.

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YEAST PRIONS ACT AS GENES COMPOSED OF SELF-PROPAGATING PROTEIN AMYLOIDS BY REED B. WICKNER, KIMBERLY L. TAYLOR, HERMAN K. EDSKES, MARIE-LISE MADDELEIN, HIROMITSU MORIYAMA, AND B. TIBOR ROBERTS Laboratory of Biochemistry and Genetics, National Institute of Diabetes, Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Genetic Criteria for Yeast Prions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Reversible Curability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Overproduction of Protien Increases the Frequency with Which the Prion Emerges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Phenotypes of the Prion Compared to Mutation of the Gene for the Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. [URE3] and URE2 Affect Nitrogen Catabolite Repression . . . . . . . . . . . . . . IV. [PSI] and SUP35 Affect Efficiency of Translation Termination. . . . . . . . . . . V. [URE3] and [PSI] as Prions of Ure2p and Sup35p, Respectively . . . . . . . . . VI. The Prion Domains of Ure2p . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Further Genetic Evidence that [URE3] is a Prion . . . . . . . . . . . . . . . . . . . . . VIII. Ure2p Is Protease Resistant in Extracts and Aggregated In Vivo in [URE3] Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX. Amyloid Formation In Vitro by Ure2p . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . X. [Het-s], a Prion of the Fungus Podospora anserina, is Necessary for a Normal Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . XI. Comparison of the Evidence for Yeast Prions with that for TSEs . . . . . . . . . XII. Implications of Yeast-Prion Amyloidoses and the Podospora Prion. . . . . . . . . XIII. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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I. INTRODUCTION The role of amyloid in mammalian spongiform encephalopathies has been controversial because only a minority of infected patients or animals show frank accumulation of amyloid as plaques, and the de novo formation of amyloid in vitro from native PrP has not been demonstrated. Nonetheless, the protease resistance of PrPSc resembles that of amyloid even when plaques are not found, and the PrP-res primed in vitro formation of PrP-res has all the properties (except for the extent of the reaction) expected for amyloid propagation (reviewed in the Chapter by Caughey et al., this volume). Unlike the rare prion diseases, amyloidoses are common features of common diseases and are often the key pathogenic factor. For most 313

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amyloidoses, there is no evidence of infectivity between individuals, although the propagation of amyloid within an individual is widely believed to occur by a seeding mechanism similar to the crystal seed mechanism proposed for prion propagation. [URE3] (Lacroute, 1971) and [PSI] (Cox, 1965) were first identified as nonchromosomal genetic elements of Saccharomyces cerevisiae, genes which did not obey Mendel’s laws, and so were thought to be due to the effects of some nucleic acid replicon that was not attached to the mitotic–meiotic segregation apparatus. The properties of such DNA plasmids, RNA viruses, the mitochondrial genome, or RNA replicons are well known; and their relationships with the host genes affecting their expression and propagation have been thoroughly described (Dujon, 1981; Wickner, 1995). It was the contrast of these nucleic acid replicons with the properties of [URE3] and [PSI] that led us to propose that the latter two elements were not the manifestation of plasmids or viruses, but were prions (Wickner, 1994). We use the term “prion” to mean “infectious protein,” not restricted to the agent causing mammalian transmissible spongiform encephalopathies (TSEs), or to any particular mechanism. II. GENETIC CRITERIA FOR YEAST PRIONS Viruses of yeast and fungi are widespread, but none exit one cell and enter another (Wickner, 1996). Whether the fungal or yeast cell wall is too great a barrier, or whether mating and hyphal anastomosis of these organisms presents too easy a method of spreading to require an extracellular route, these viruses are spread only by cell-cell fusion. They are often first found as nonmendelian genetic elements or, if there is no phenotype associated with the replicon, as nucleic acids whose presence shows nonmendelian inheritance. One would thus expect a yeast prion to also appear as a nonmendelian element. [Note that the terms nonmendelian (genetic) element, nonchromosomal genetic element, and cytoplasmic genetic element are essentially synonymous; see Table I.] We use the word prion to mean an infectious protein, a protein that is altered in some way so that it does not carry out its normal function properly, but has acquired the ability to convert the normal form of the protein to the same abnormal (nonfunctional) form. By this definition, three properties are expected of a prion that are in contrast to the corresponding properties of a nucleic acid replicon (Wickner, 1994). A. Reversible Curability If a yeast virus or plasmid (or the mitochodrial DNA genome) is cured from a strain, they will not return or arise again unless they are

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TABLE I Properties that Distinguish Chromosomal Genes from Nonchromosomal Genes Property Induced loss of the gene Mitotic segregation of alleles Segregation of alleles in meiosis Transfer by cytoplasmic transfer

Nonchromosomal Gene

Chromosomal Gene

Efficient curing by nonmutagenic agents High frequency 4+:0– or irregular Efficient

Induced mutation is still a rare event Low frequency 2+:2– Rare

introduced from another cell. In contrast, the curing of a prion cannot be irreversible, because the normal form of the protein is still present and can again undergo the same spontaneous change (whatever that may be) to form the prion (Wickner, 1994). This de novo change is likely to be a rare event, but not as rare as the appearance of a new virus. B. Overproduction of the Protein Increases the Frequency with Which the Prion Emerges Plasmids and viruses often depend heavily on chromosomally encoded proteins, but overproduction of such proteins will not induce the viruses to arise de novo. In contrast, overproduction of a protein that has the potential to undergo a prion change should increase the frequency with which that change is observed, simply because there is more of the protein present to change (Wickner, 1994). This effect may be amplified in the case of [URE3], [PSI], and scrapie (and all amyloidoses) because the crystal seed priming mechanism predicts that the frequency of primary nidus formation will increase as a power of the concentration of monomer (Lansbury and Caughey, 1995). C. Phenotypes of the Prion Compared with Mutation of the Gene for the Protein Mutation of a chromosomal gene needed for propagation of a virus or plasmid results in the opposite phenotype from that owing to the presence of the plasmid (see Table II). A prion depends for its propagation on the continued production of the protein from the chromosomal gene that encodes it. If the prion produces a phenotype by depleting the amount of the normal form, it will produce the same phenotype as does mutation in the chromosomal gene encoding the protein (Wickner, 1994) (Table II). As described later, the properties of the nonmendelian genetic elements, [URE3] and [PSI], in relation to these genetic criteria, indicated

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proline media, it would readily substitute for uracil allowing growth of the ura2 mutants. Mutants able to take up USA in the presence of ammonia defined the chromosomal ure1 and ure2 genes, and a nonchromosomal genetic element (gene) that he named [URE3] (Drillien et al., 1973; Drillien and Lacroute, 1972; Lacroute, 1971; Schoun and Lacroute, 1969b). Why is USA uptake regulated by the nitrogen source? USA is very similar to allantoate, a poor but usable nitrogen source for yeast, whose uptake is carried out by Dal5p, and the DAL5 gene is regulated by the nitrogen catabolite repression pathway (Rai et al., 1987; Turoscy and Cooper, 1987) (Fig. 1). Ure2p is a key component in nitrogen catabolite repression, receiving signals from Mks1p (Edskes et al., 1999b) and inhibiting the action of the positively acting “GATA” transcription factor Gln3p (Coschigano and Magasanik,

FIG. 1. The role of Ure2p in regulation of nitrogen catabolism. Ure2p is part of a signal transduction pathway by which the presence of a good nitrogen source (ammonia) represses production of proteins involved in utilization of poor nitrogen sources, such as allantoate (Coschigano and Magasanik, 1991; Courchesne and Magasanik, 1988; Edskes et al., 1999b; Mitchell and Magasanik, 1984)(see text). The coincidental resemblance of ureidosuccinate (an intermediate in uracil biosynthesis) and allantoate result in uptake of ureidosuccinate by Dal5p, the allantoate permease (Rai et al., 1987; Turoscy and Cooper, 1987). Because DAL5 is under Ure2p control, ureidosuccinate uptake measures Ure2p activity.

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1991; Courchesne and Magasanik, 1988; Mitchell and Magasanik, 1984). Gln3p is necessary for the transcription of DAL5 and many other genes needed for the utililization of poor nitrogen sources. Thus, USA uptake (by Dal5p) is regulated by nitrogen catabolite repression (Fig. 1), thus explaining the identification of ure2 and [URE3] in the genetic screen by Lacroute. IV. [PSI] AND SUP35 AFFECT EFFICIENCY OF TRANSLATION TERMINATION Sup35p and Sup45p are the subunits of the translation release or termination factor (Fig. 2). They form a heterodimer that recognizes the termination codon and cleaves the completed peptide from the final tRNA, thereby terminating the translation process. All termination events involve a competition between the normal action of the Sup35p/Sup45p complex and the misrecognition of the termination codon by a tRNA to result in readthrough and continuation of the peptide chain. Nonsense suppressor tRNAs have an altered anticodon, which instead of recognizing the codon for the corresponding amino acid now recognizes one or more termination codons. Both sup35 and sup45 mutations, by impairing the normal termination process, make weak suppressor tRNAs strong and strong suppressor tRNAs lethal. In 1965, Cox discovered a nonmendelian genetic element that had the same effect as a sup35 or sup45 mutation (Cox, 1965). He named this element [PSI].

FIG. 2. Sup35p and Sup45p constitute the translation release (termination) factor (Stansfield et al., 1995; Zhouravleva et al., 1995). Thus, mutation of SUP35 or conversion of Sup35p to a prion increases the efficiency of weak nonsense-suppressing tRNAs.

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V. [URE3] AND [PSI] AS PRIONS OF URE2P AND SUP35P, RESPECTIVELY [URE3] can be cured by growth of cells in media containing 1 mM guanidine (M. Aigle, cited in [Cox et al., 1988]), but this curing is reversible in that from the cured strains can again be isolated [URE3] derivatives (Wickner, 1994). Overproduction of Ure2p results in a 20- to 200-fold increase in the frequency with which [URE3] arises de novo (Wickner, 1994). Finally, the phenotype of [URE3] strains and ure2 mutants are essentially the same (Lacroute, 1971), and URE2 is necessary for the propagation of [URE3] (Aigle and Lacroute, 1975; Wickner, 1994). Thus, [URE3] has the properties expected of a prion of Ure2p (Wickner, 1994) (Fig. 3). [PSI] can be cured by growth of cells in high osmotic strength media (Singh et al., 1979) or by 1 mM guanidine-HCl (Tuite et al., 1981), but in each case, curing is reversible in that clones that have acquired [PSI] may again be selected from the cured strain (Chernoff et al., 1993; Lund and Cox, 1981). Overproduction of Sup35p results in [PSI] arising de novo at 100 times the normal frequency (Chernoff et al., 1993; Derkatch et al., 1996). Finally, the phenotype of [PSI] is essentially the same as that of a chromosomal sup35 mutant, yet SUP35 is necessary for the propagation of [PSI] (Doel et al., 1994; TerAvanesyan et al., 1994). For these reasons, it was proposed that [PSI] is a prion of Sup35p (Wickner, 1994). VI. THE PRION DOMAINS OF URE2P Deletion analysis of URE2 showed that the N-terminal 65 residues were dispensable for its nitrogen regulation function (Coschigano and Magasanik, 1991; Masison and Wickner, 1995), but this region, when overexpressed, was quite active in inducing the de novo appearance of [URE3] (Masison and Wickner, 1995). A molecule lacking the 1–65 region was, in fact, unaffected by introduction of [URE3] into the cell (Masison et al., 1997; Masison and Wickner, 1995). Furthermore, cells expressing Ure2p1–65 in the complete absence of the C-terminal domain were capable of propagating [URE3] (Masison et al., 1997). Thus, Ure2p1–65 was designated the prion domain and Ure2p66–354 was named the nitrogen regulation domain (Masison and Wickner, 1995). Further work showed that residues 1–80 were an even more efficient inducer of [URE3] appearance than 1–65 (Maddelein and Wickner, 1999) and that other parts of the Ure2 molecule could influence the efficiency of this process. Deletion of residues 221–227 results in inability of

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FIG. 3. Genetic criteria for a prion illustrated by [URE3] (Wickner, 1994) (see text).

the otherwise intact overproduced Ure2p to induce [URE3] (Maddelein and Wickner, 1999). Deletion of residues 151–157 or 348–354, both parts of the nitrogen regulation domain, result in elevated prion-inducing ability by the rest of the molecule (Maddelein and Wickner, 1999), and the 1–65 or 1–80 regions are hundreds or thousands of times better inducers than the full-length molecule. We interpret these results to mean that the C-terminal nitrogen regulation domain interacts with the N-terminal prion domain and stabilizes it in the normal form. The prion-promoting and prion-inhibiting regions of Ure2p are summarized in Fig. 4. Remarkably, two nonoverlapping prion-inducing fragments can be made from Ure2p. Although the Ure2p66–354 region is inactive in induc-

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FIG. 4. Prion—promoting and prion—inhibiting regions of Ure2p.

ing [URE3], further deletion of residues 151–157 and 348–354 (two prion-inhibiting regions) makes this fragment able to induce [URE3] (Maddelein and Wickner, 1999). This activity is dependent on the asparagine-rich residues 66–80 and the 221–227 region. The Ure2 prion domain is quite rich in asparagine, and deletion of runs of asparagine makes the otherwise intact Ure2p molecule nearly inactive in inducing [URE3] (Maddelein and Wickner, 1999). It is possible that the role of asparagine runs in [URE3] is related to the role of runs of glutamine in Huntington’s disease (Stott et al., 1995). Native Ure2p isolated from yeast is a dimer (Taylor et al., 1999), as is that produced in Escherichia coli (Perrett et al., 1999). Chemical denaturation studies of a series of deletion mutants showed that the absence of the prion domain did not affect the overall stability of the Ure2p molecule (Perrett et al., 1999). This indicates that any energy of interaction of the C-terminal and N-terminal domains is equivalent to that of alternative C-terminal – C-terminal interations. VII. FURTHER GENETIC EVIDENCE THAT [URE3] IS A PRION Because Ure2p is a transcription regulatory factor, it was particularly important to eliminate the possibility that [URE3] was not a prion, but a metastable regulatory state based on, for example, a positive feedback transcription regulation loop. The fact, discussed previously, that prion domains and nitrogen regulatory regions of Ure2p could be distinguished already indicated that [URE3] was not such a regulatory loop. It was further shown that [URE3] could be propagated in cells with either repressed or derepressed nitrogen regulation (Masison et al., 1997). Moreover, the nitrogen regulatory state did not have a detectable

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effect on induction of [URE3] by the overproduction of Ure2p (Masison et al., 1997). It was critical to show that the induction of [URE3] appearance was really de novo. If [URE3] were a mutant form of a putative wild-type plasmid or virus that is dependent on Ure2p for its propagation, one could explain the [URE3] phenotype if, for example, [URE3], like the defective interfering viruses or “suppressive petite” mitDNA mutants, eliminated the normal genome by competing with it. Then a ure2∆ strain would lack the putative wild-type plasmid, and in such a strain, the infectious [URE3] could not be produced. Into such a ure2∆ mutant was introduced a Ure2p-overproducing plasmid. When Ure2p production was turned on, [URE3] clones arose as usual. These [URE3]s were shown to be infectious, curable, and dependent on continued Ure2p production for their propagation (Masison et al., 1997). This showed that [URE3] really could arise de novo. “Spontaneous generation” can happen for prions. It was shown that induction of [URE3] was really occurring by overproduction of the Ure2 protein and not by overproduction of the URE2 mRNA or the presence of the gene itself in high copy number. This was done by showing that the reading frame of the prion domain was critical for induction of [URE3] appearance, and that the parts of the mRNA changed to affect the reading frame were not critical (Masison et al., 1997). Nor was the level of mRNA affected by these changes. Of course, the gene in high copy number, if not transcriptionally activated, did not induce the appearance of [URE3]. VIII. URE2P IS PROTEASE RESISTANT IN EXTRACTS AND AGGREGATED IN VIVO IN [URE3] CELLS The first confirmation that Ure2p was altered in [URE3] cells was the finding that it was relatively protease-resistant in extracts of such strains compared with isogenic wild-type extracts (Masison and Wickner, 1995). Normal Ure2p is rapidly digested, but the Ure2p in the extracts of [URE3] cells is converted first to two relatively resistant fragments of 30 and 32 kDa. In some [URE3] strains, there is a 7 to 10 kDa species that is stable throughout the digestion period. The location of Ure2p can be determined using fusions of Ure2p with the green fluorescent protein (GFP) (Edskes et al., 1999a). The fusion protein is active in carrying out nitrogen regulation and is inactivated by the introduction into the cell of the [URE3] prion. In [ure-o] strains, the Ure2p-GFP fusion protein is found to be evenly distributed

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in the cell, except perhaps in the vacuole and nucleus. In [URE3] strains, the fusion protein is aggregated (Fig. 5, see color insert). Curing [URE3] from the strain by growth on guanidine results in the fusion protein again showing the even distribution characteristic of the wild-type cells. A fusion protein with just the prion domain (residues 1–65) of Ure2p with GFP shows similar aggregation in [URE3] cells and an even distribution in wild-type or cured strains (Edskes et al., 1999). In contrast, a fusion with the C-terminal nitrogen regulation domain shows no evidence of aggregation in [URE3] cells, as expected from its lack of inactivation by introduction of the prion (see previously). These results indicate that Ure2p is in an aggregated state in [URE3] strains that makes Ure2p partially protease resistant. IX. AMYLOID FORMATION IN VITRO BY URE2P Amyloid is an abnormal filamentous form of protein characterized by high β-sheet content, protease-resistance, and yellow-green bi-refringence on staining with Congo red (Kisilevsky and Fraser, 1997). Amyloid is a prominent feature of (among others) Alzheimer’s disease, late-onset diabetes, persistent acute inflammatory conditions, multiple myeloma, and transmissible spongiform encephalopathies. The chemically synthesized prion domain peptide, Ure2p1–65, was found to rapidly and spontaneously form filaments (Fig. 6A) when diluted from 6 M guanidine into a neutral buffer (Taylor et al., 1999). These filaments are about 50 Å in diameter, are resistant to digestion by proteinase K, are almost entirely β sheet in structure, and show the yellow-green bi-refringence on staining with Congo red that is typical of amyloid (Fig. 7, see color insert). In vivo, the prion domain induces the prion formation by the fulllength Ure2 protein. Dilution of Ure2p1–65 into a solution of the fulllength native soluble Ure2p results in the formation of “co-filaments” (Fig. 6B) (Taylor et al., 1999). An equimolar amount of Ure2p1–65 results in complete incorporation of native Ure2p into the insoluble filaments under conditions in which the native Ure2p is stably soluble. These cofilaments are 200 Å in diameter and require about 24 hours to form. Like the filaments formed from the prion domain alone, the cofilaments have all of the properties of amyloid. They are high in βsheet content, and calculation indicates that the full-length molecules have increased their β-sheet content, as have the prion domain fragments. The cofilaments also show yellow-green bi-refringence on staining with Congo red. Whereas the native Ure2p is rapidly digested,

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treatment of the cofilaments with proteinase K produces partially resistant 30 and 32 kDa fragments and a strongly resistant 7 to 10 kDa fragment that is derived from the prion domain (Taylor et al., 1999). This pattern is closely similar to that seen in comparing extracts of wild-type and [URE3] strains (Masison and Wickner, 1995). The cofilament formation is a highly specific process. Dilution of Ure2p1–65 into a mixture of proteins results in only full-length Ure2p incorporation into the filaments formed. Likewise, filament formation by the Aβ peptide in the presence of native Ure2p does not result in incorporation of the Ure2p into the filaments formed (Taylor et al.,

FIG. 6. Amyloid filaments formation by native Ure2p is induced by the prion domain, Ure2p1–65 (Taylor et al., 1999). (A) Filaments 50 Å in diameter formed by the prion domain peptide, Ure2p1–65. (B) A 200 Å diameter “cofilaments” formed by an equimolar mixture of Ure2p1–65 and native full length Ure2p. (C) Treatment of cofilaments from (B) with proteinase K leaves narrow filaments composed of the prion domain peptide and the prion domain of Ure2p. (D) Cofilament seed formation of 400 Å diameter filaments by excess native Ure2p.

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1999). This is the specificity expected of the in vivo [URE3] phenomenon, which, like other amyloidoses, is specific for the protein affected. Addition of a small amount of the cofilaments to a 40-fold excess of native Ure2p induces filament formation by the native Ure2p over a period of days to weeks (Fig. 6D) (Taylor et al., 1999). These “seeded filaments” are about 400 Å in diameter and show the same protease-resistance and Congo red bi-refringence as do the cofilaments. Thus, the amyloid formation seems capable of indefinite propagation once it has been initiated by the prion domain peptide. The properties of amyloid formation in vitro initiated by the Ure2p prion domain peptide shows all the properties expected if self-propagating amyloid is the explanation of the [URE3] phenomenon in vivo. The initiation of amyloid formation in vitro by the prion domain parallels its initiation of prion formation in vivo. The aggregation of Ure2p in [URE3] cells parallels its aggregation in the form of amyloid in vitro. Finally, the pattern of protease resistance of Ure2p in extracts of [URE3] cells is the same as that seen in amyloid formed in vitro. X. [HET-S], A PRION OF THE FUNGUS PODOSPORA ANSERINA, IS NECESSARY FOR A NORMAL FUNCTION Filamentous fungi have two modes of cell-cell fusion. The purpose of sexual fusion is to generate diversity through meiosis. This function is reflected in the genetic control of mating type: Only strains of different mating type can mate to form diploids. In contrast, the purpose of hyphal anastomosis (fusion of the cell processes of two colonies) is to share nutrients. However, hyphal anastomosis carries with it the danger of transmission of a virus present in one strain to the other strain. Thus, fungi restrict the partners with whom they will engage in hyphal anastomosis to those that are apparently genetically identical and thus already have the same viruses. Identity is assured by requiring identity at a series of chromosomal loci, called het loci. When two incompatible strains meet, a few peripheral hyphae fuse, but these fused hyphae die and somehow form a barrier to further fusion between the colonies (Begueret et al., 1994). One such locus in Podospora is the het-s locus, with alleles het-s and hetS. Hyphae of a het-s strain can fuse with another het-s strain, but not with a het-S strain (Fig. 8). The het-s locus encodes a 289 residue protein with no similarity to other proteins in the database (Turcq et al., 1991; Turcq et al., 1990). The difference between het-s and het-S is 14 amino acids, with the residue 33 difference critical to produce the heterokaryon incompatibility reaction (Deleu et al., 1993).

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Rizet found that genotypically het-s cells could have either of two phenotypes: they could be incompatible with het-S strains as described previously, or they could be neutral, fusing readily with either het-s or het-S colonies (Rizet, 1952). He found that this difference was controlled by a nonchromosomal genetic element, called [Het-s] (Fig. 8) (BeissonSchecroun, 1962; Rizet, 1952). Its absence is denoted [Het-s*]. The hets [Het-s] strains showed the expected incompatibility with het-S cells, but het-s [Het-s*] strains were neutral. [Het-s] is lost almost uniformly from the offspring of a cross of the type [Het-s] X [Het-s*]; this is a form of curing. However, from the meiotic segregants lacking [Het-s] again arise cells carrying [Het-s] (Beisson-Schecroun, 1962). This is reversible curing, a property of a prion. The propagation of [Het-s] requires the het-s product, and overexpression of the het-s protein results in an increased frequency with which [Het-s] arises (Coustou et al., 1997). Here again are the genetic properties expected of a prion. In addition, the het-s encoded protein is relatively protease-resistant in extracts of [Het-s] strains compared with extracts of wild-type strains (Coustou et al., 1997). This led Coustou et al. to suggest that [Het-s] is a prion form of the protein encoded by the het-s gene (Coustou et al., 1997). [Het-s] is the first clear case of a prion that is necessary for a normal cellular function. Heterokaryon incompatibility does involve some cell

FIG. 8. The [Het-s] prion of Podospora anserina is necessary for heterokaryon incompatibility, a normal cellular function (Coustou et al., 1997).

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death, but, like apoptosis, it seems to be death with a definite purpose. It is a general feature of filamentous fungi, not an isolated disease state. In other fungi and with other het loci, there is no known involvement of nonmendelian genetic elements. This work raises the issue of what other normal cellular functions could be mediated by prion-like mechanisms. Sonneborn discovered that the pattern of cilia on the cuticle of Paramecium was a self-propagating trait (Beisson and Sonneborn, 1965). Alteration of the pattern by an accident of mating or by surgery produced an altered self-propagating pattern. This finding has been extended to other organisms (Grimes and Aufderheide, 1991). It is possible that other cellular structures are not self-assembling, but are self-patterning. Such a structure might show up genetically as a mutant that could not assemble the structure, but whose phenotype was not restored by introduction of the normal gene by transformation. Peroxisomes were believed to be such a structure, but such mutants have not been found despite intensive searches (reviewed in Lazarow and Kunau, 1997). XI. COMPARISON OF THE EVIDENCE FOR YEAST PRIONS WITH THAT FOR TSES Table III compares the bases for the prion model for scrapie with the yeast and fungal systems. The exceptional UV-resistance of the scrapie agent (Alper et al., 1967) provided the initial evidence and prompted the initial formulation of the concept that the scrapie agent was composed of only protein (Griffith, 1967). The early studies of Dickinson first identified PrP as the Sinc gene (Dickinson et al., 1968), and purification of the infectious agent suggested that this protein might be the scrapie infectious agent (Prusiner, 1982). However, the best preparations have 105 molecules of PrP, suggesting there could be another critical molecule or modification of PrP. Lacking an external transfection system, the yeast systems do not have this line of evidence available, but it has been shown that overproduction of the Ure2 and Sup35 proteins (not the genes in high copy number or the mRNAs) make [URE3] and [PSI] arise de novo (Derkatch et al., 1996; Masison et al., 1997), a result at once cleaner and in some sense equivalent to the PrP purification result. We refer to this as the “molecular biologic purification” experiment. The scrapie system has not yet been shown to satisfy any of the three genetic criteria. Scrapie cannot be cured, so reversible curing cannot yet be tested. Overproduction of PrP does produce a disease in mice,

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TABLE III Comparison of Evidence for the Putative Prion Systems

Genetic evidence Reversible curing Overproduction → ↑ prion de novo Phenotype relationship Alleles control infection Epidemiology: spontaneous, inherited and infectious disease Biochemical evidence UV-resistance of infectivity Biochemical purification of the agent Molecular biological purification of the agent In vitro system a

TSEs

[URE3]

[PSI+]

[Het-s]

–a – – + +

+ + + + +

+ + + + +

+ + – + +

+ + –

– – +

– – +

– – –

+

+

+

–

–, not yet proved.

but no infectious material (Westaway et al., 1994); thus it is not scrapie that is being observed. The phenotypic relationship is not available in the case of scrapie because PrPSc evidently positively harms the organism; the scrapie disease is not produced by absence of PrP (Bueler et al., 1992). That PrP is necessary for scrapie (Bueler et al., 1993) shows that PrP is important, but is not evidence that it is itself the scrapie agent. The control of the infection specificity by the PrP sequence (Prusiner et al., 1990) again shows PrP to be central to the disease process. The in vitro systems for scrapie (Bessen et al., 1995; Kocisko et al., 1994; Kocisko et al., 1995), [URE3] (Taylor et al., 1999) and [PSI] (Glover et al., 1997; King et al., 1997; Paushkin et al., 1997) all seem to faithfully reproduce the in vivo features, strongly suggesting that they are reproducing the infectious processes. The presence of PrP amyloid in TSE brain has been well documented, but there remains disagreement as to whether amyloid is central to the infectious or pathogenic processes. XII. IMPLICATIONS OF YEAST PRION-AMYLOIDOSES AND THE PODOSPORA PRION As new types of amyloid are discovered—many of them very common and apparently associated with disease (Haggqvist et al., 1999; Wester-

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mark et al., 1995)—the importance of a thorough investigation of the genetic mechanisms governing cellular handling of amyloid increases. We expect that yeast amyloidoses will be useful in this regard. The finding that the [Het-s] prion of the filamentous fungus Podospora is essential for for a normal function suggests that other normal functions will be found to involve a prion mechanism. XIII. SUMMARY We proposed genetic properties by which prions can be recognized among infectious elements. This indicated that [URE3] and [PSI], two nonchromosomal genetic elements of yeast, were prions (infectious altered forms) of Ure2p and Sup35p, respectively. Overexpression of Ure2p leads to the de novo appearance of the [URE3] prion, and the N-terminal 65–80 residues (the prion domain) are specifically responsible for this prion-inducing activity. The prion domain is sufficient to propagate [URE3] and is necessary for a Ure2p molecule to be affected by [URE3]. The remaining C-terminal residues 81–354 are responsible for nitrogen catabolite repression, the normal function of Ure2p. Ure2p is proteaseresistant specifically in extracts of [URE3] strains and is aggregated in vivo specifically in such strains. The chemically synthesized Ure2p prion domain (Ure2p1–65) spontaneously forms classic amyloid filaments (50 Å diameter) in vitro, and specifically induces the native full length Ure2p to form a 1:1 amyloid cofilament (200 Å). These amyloid cofilaments can prime amyoid filament formation by an excess of native Ure2p. The features of the in vitro amyloid propagation reaction appear to reproduce the in vivo properties of [URE3] prion propagation. This system may be useful for detecting new prions, finding amyloid-inducing and amyloid-curing agents, and determining the cellular components that affect the initiation and propagation of infectious amyloids. The [Het-s] prion was found in the filamentous fungus Podospora anserina by similar genetic tests to those we used for [URE3] and [PSI]. [Het-s] is necessary for a normal function of Podospora cells, heterokaryon incompatibility. This suggests that other normal cellular functions may involve a prion-like mechanism. NOTE ADDED IN PROOF Recently, the presence of amyloid filaments of Ure2p in cells carrying the [URE3] prion has been demonstrated by immuno-electron microscropy (Speransky et al., 2001). The filaments were observed to form

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large networks in the cytoplasm, and were never observed in [ure-o] cells. The same study showed that much of the Ure2p in extracts of [URE3] strains is in a form insoluble in boiling 2% SDS and 3 M urea, consistent with amyloid structure. The de novo generation of [URE3] is now known to require the Mks1 protein, although propagation of [URE3] is not affected by the absence of Mks1p (Edskes and Wickner, 2000). Mks1p was first defined as a protein whose overexpression slows cell growth, and this growthinhibiting activity was found to be antagonized by the Ras-cAMP pathway (Matsuura and Anraku, 1993), and then found to be part of the nitrogen catabolite repression pathway (Edskes, et al., 1999). Introduction of a constitutive (“oncogenic”) RAS2 allele (Ras2val19) also blocks de novo generation of the [URE3] prion, as predicted from the known action of Ras-cAMP on Mks1p (Edskes and Wickner, 2000). This implies that signal transduction pathways can affect prion generation. Propagation of [PSI] is known to require an optimal level of the chaperone Hsp104 (Chernoff et al., 1995), with loss of [PSI] observed by either deletion or overexpression of HSP104. Recently, it was shown that deletion of HSP104 also results in the loss of [URE3], but that overexpression of HSP104 does not affect [URE3] propagation (Moriyama et al., 2000). However, overexpression of Ydj1p, a member of the Hsp40 family (Lu and Cyr, 1998), does cure [URE3] (Moriyama et al., 2000). These results suggest that many (perhaps all) prions are affected by chaperones, but the details may depend on the detailed properties of the individual amyloid structures. The structure of the nitrogen regulation domain of Ure2p has been determined recently by X-ray crystallography (Bousset et al., 2001; Umland et al., 2001). The structure closely parallels that of glutathione S-transferases, as expected from the amino acid sequence homology of Ure2p to many such enzymes (Coschigano and Magasanik, 1991).

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[PSI +], SUP35, AND CHAPERONES TRICIA R. SERIO* AND SUSAN L. LINDQUIST† * Department of Molecular Genetics and Cell Biology, and † The Howard Hughes Medical Institute, The University of Chicago, Chicago, Illinois 60637

I. [PSI +] . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Formulation of the Prion Hypothesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Genetic and Cell Biological Support for [PSI +] as a Yeast Prion . . . . . . . . . . A. Link to SUP35 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. PSI + Induction by Over-expression of Sup35. . . . . . . . . . . . . . . . . . . . . . . C. Alternative Phenotypes Associated with Distinct Protein States. . . . . . . . D. Replication of the Alternate Physical State . . . . . . . . . . . . . . . . . . . . . . . . IV. A Model for the [PSI +] Phenotype . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Crucial Residues in the Replication of Protein States . . . . . . . . . . . . . . . . . . A. Sup35 Sequence Elements (N, M, C) . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Two Regions in N Contribute to PSI + Inheritance . . . . . . . . . . . . . . . . . . VI. Modeling [PSI +] In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Sup35 Self-assembly In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Replication of the Alternate Protein State In Vitro . . . . . . . . . . . . . . . . . . C. Biochemical Effects of N Mutations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Regulation of [PSI] Metabolism by Molecular Chaperones . . . . . . . . . . . . . A. Hsp104 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Ssa1. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Ssb . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

335 336 338 338 339 339 339 341 342 342 344 346 346 348 351 353 354 360 361 362 363

I. [PSI +] In 1965, Cox described an unusual genetic element, [PSI +], that modified translation termination efficiency in the yeast Saccharomyces cerevisiae (Cox, 1965). Evidence gathered for more than 30 years by genetic analyses, combined with the results of more recent cell biological, molecular genetic, and biochemical experiments, presents an overwhelmingly convincing case that this genetic element is proteinacious. Specifically, a change in phenotype is brought about by a self-perpetuating change in the conformation of a particular protein, rather than by a change in a nucleic acid. Remarkably, the process appears to have many features in common with certain protein misfolding diseases that are discussed elsewhere in this volume. Moreover, several other proteins in yeast are capable of self-perpetuating changes in conformation that allow them to serve as genetic elements, producing heritable changes in phenotype. One of these, [URE3], is the subject of another 335 ADVANCES IN PROTEIN CHEMISTRY, Vol. 57

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article in this volume. Here, we focus on [PSI +] and recent progress in understanding its conformational transitions. II. FORMULATION OF THE PRION HYPOTHESIS Early characterization of the [PSI +] determinant was based on its pattern of inheritance in genetic crosses. In these experiments, transmission of the [PSI +] determinant was monitored through inheritance of its associated phenotype. In [psi–] strains, translation terminates faithfully at all stop codons, but in [PSI +] strains, stop codons are readthrough at a low frequency (Cox, 1965). This difference in translation termination is detectable in strains carrying stop mutations in various genetic markers because read-through suppresses the effects of the mutation. The [PSI +] element is inherited in an orderly, reproducible way but one that is different from most genetic traits (Fig. 1) (Cox, 1965). When a haploid [PSI +] strain is mated to a haploid [psi –] strain, the resulting diploid has a suppression phenotype; that is, [PSI +] is dominant. On sporulation, however, none of the haploid progeny are [psi–] as would be expected for a nuclear determinant. Instead, [PSI +] is transmitted to all haploid progeny. (The capital letters in [PSI +] signify dominance; the brackets signify nonchromosomal inheritance.) This unusual pattern of inheritance was partly explained by later experiments that localized the [PSI +] determinant to the cytoplasm (Cox et al., 1980; Fink and Conde, 1976). Surprisingly, however, the [PSI +] phenotype could not be linked to any of the known cytoplasmic nucleic acids in yeast (Cox et al., 1988; Serio and Lindquist, 1999). [PSI ] is stably inherited in both normal mitotic divisions and in sexual meiotic divisions (Cox, 1965). However, conversion from the [PSI +] to the [psi –] state, known as [PSI +] curing, occurs spontaneously at a low frequency (Cox, 1965; Lund and Cox 1981). Curing can be induced by treatment with several agents that are not mutagenic to nucleic acids. For example, [PSI +] strains grown in the presence of methanol (10% v/v) or guanidine HCl (1 to 5 mM) are cured quantitatively (Eaglestone et al., 2000; Tuite et al., 1981). Conversely in [psi –] strains, [PSI +] appears de novo at a higher rate if cells carry extra copies of the SUP35 gene (Chernoff et al., 1992a). Remarkably, these strains remain [PSI +] when the extra copies of Sup35 are lost (Chernoff et al., 1993). In the absence of spontaneous generation, this reappearance of [PSI +] is difficult to reconcile with a nucleic acid determinant (Cox et al., 1988; Serio and Lindquist, 1999).

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FIG.1. [PSI +] is cytoplasmically inherited. Black circles: [PSI +] determinant. Yeast cells can grow either as haploids (one copy of each chromosome) or diploids (two copies of each chromosome). When a [PSI +] haploid is mated to a [psi –] haploid, the resulting diploid is [PSI +], indicating that [PSI +] is dominant. On sporulation, each nuclear determinant (gray fill and wavy lines) segregates to half the haploid progeny (2:2), and the cytoplasmic [PSI +] determinant segregates to all progeny (4:0).

In 1994, Wickner proposed that both [PSI +] and another cytoplasmic determinant in yeast, [URE3], are inherited by a previously unrecognized mechanism that is consistent with the unusual genetic features of each. He suggested that the [PSI +] and [URE3] phenotypes were each transmitted by changes in the physical states of the Sup35 or Ure2 proteins, respectively, rather than by an alteration in a nucleic acid (Wickner 1994). That is, the [PSI +] and [URE3] determinants were hypothesized to be proteins rather than nucleic acids. This revolutionary concept provides a plausible explanation for the reversible curing of both [PSI +] and [URE3]; a switch in the physical state of a protein already present in the cell is all that is required to induce a heritable change in phenotype.

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This concept, that proteins could act as elements of genetic inheritance, was based on the mammalian prion hypothesis. This hypothesis had been previously proposed to explain the unusual nature of the infectious agent responsible for a group of devastating neurodegenerative disorders in mammals, collectively known as the transmissible spongiform encephalopathies (TSEs) (Griffith, 1967; Prusiner, 1982). The hypothesis predicts that a single protein can stably exist in at least two alternative physical states, each of which is associated with a distinct phenotype. (For the mammalian prion protein, PrP, that phenotype is a progressive pathology.) Normally, the prion protein will exist in one of these states. The altered prion form spontaneously arises at an extremely low frequency. Once the prion form appears, it shifts the equilibrium in its favor by influencing protein in the normal state to adopt the prion form. This self-perpetuation of physical states “replicates” the information that specifies a change in phenotype and is thereby analogous to the replication of nucleic acid determinants. It is the cornerstone of the hypothesis that proteins can serve as genetic elements. III. GENETIC AND CELL BIOLOGICAL SUPPORT FOR [PSI +] AS A YEAST PRION The yeast prion hypothesis provides explanations for several unique characteristics of [PSI +] that were inconsistent with a nucleic acid determinant (Wickner, 1994). A. Link to SUP35 Although the phenotype of a prion is predicted to be inherited through the transmission of an alternative protein state, it will also be dependent on the gene that encodes that protein. Expression of the gene provides a supply of protein that can then be converted to the prion state. In the absence of new protein, the associated phenotype will be lost because the determinant (a particular structural state of the protein) cannot be replicated. [PSI +] exhibits such a relationship with the protein Sup35. Sup35 is an essential protein, and its gene cannot be deleted. However, deletion of the N-terminus of SUP35 or mutations at certain residues in this region (described in detail later) lead to the irreversible loss of [PSI +] (Doel et al., 1994; Liu and Lindquist, 1999; McCready et al., 1977; Ter-Avanesyan et al., 1994; Young and Cox, 1971). Such mutations are referred to as [PSI +]-no-more (pnm) (Young and Cox, 1971).

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B. [PSI +] Induction by Overexpression of Sup35 The prion hypothesis predicts that the spontaneous appearance of the prion state and its associated phenotype is a rare and stochastic event. Increasing prion protein expression levels should lead to an increase in the rate at which this event occurs (Prusiner, 1982). [PSI +] is readily induced in [psi –] strains by episomal plasmids encoding the SUP35 gene (Chernoff et al., 1993; Chernoff et al., 1992a). If a nonsense mutation is placed at the beginning of the SUP35 open reading frame (ORF) in these plasmids, the ability to induce [PSI +] de novo is lost (Derkatch et al., 1996). This observation suggests that it is the Sup35 protein rather than the DNA or the messenger RNA that is active in the process. C. Alternative Phenotypes Associated with Distinct Protein States The yeast prion hypothesis holds that distinct physical states of a single protein form the basis of alternative phenotypes (Wickner, 1994). Indeed, when yeast lysates are subjected to high-speed centrifugation, Sup35 is found mostly in the soluble fraction in [psi–] strains, but in [PSI +] strains, Sup35 is largely insoluble (Fig. 2A) (Patino et al., 1996; Paushkin et al., 1996). In addition, the Sup35 protein of [PSI +] cells has increased resistance to proteolysis compared with that of [psi –] cells (Patino et al., 1996; Paushkin et al., 1996). These differences in physical state are tightly linked to the [PSI +] phenotype; in sequential rounds of curing and de novo induction of [PSI +], Sup35p changes from insoluble→ soluble→insoluble (Patino et al., 1996; Paushkin et al., 1996). Finally, transient changes in the levels of the molecular chaperone Hsp104 induce a heritable change in phenotype from [PSI +] to [psi –] and a corresponding change in Sup35p from insoluble→soluble (discussed in detail later) (Chernoff et al., 1995; Patino et al., 1996). Because the only known function of Hsp104 is to alter the physical state of other proteins (see later), this observation also provides a strong genetic argument for a protein-only mode of inheritance for [PSI +]. D. Replication of the Alternate Physical State The prion hypothesis predicts that the prion conformation replicates by influencing new protein to adopt the same state (Griffith, 1967; Prusiner, 1982). A series of experiments using fusions of Sup35 to the green fluorescent protein (GFP) have directly demonstrated this process in the case of [PSI +] (Fig. 2B) (Patino et al., 1996). Fluorescence

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FIG.2. Sup35 adopts an alternate state in the [PSI +] cytoplasm. (A). Differential centrifugation. Lysates of [psi –] and [PSI +] strains were subjected to centrifugation at 12,000g and the supernatant (S) and pellet (P) fractions were analyzed by SDS-PAGE and immunoblotting. Sup35 is mostly soluble in [psi –] lysates, while [PSI +] is found largely in the pellet fraction in [PSI +] lysates. The distribution of the ribosomal protein L3 is unaltered by the presence of [PSI +]. (The small quantity of pelletable material in [PSI –] cells is not due to protein in the [PSI +] state but to denaturation or association with ribosomes.) (B) Fusion protein fluorescence. Fluorescence from a Sup35 fusion to the green fluorescent protein (GFP) is diffusely distributed throughout the [psi –] cytoplasm when expressed for a brief period (4 hours). Fluorescence from the same fusion protein rapidly coalesces into discrete foci within the same time frame in [PSI +] cells because it is captured by preexisting Sup35 complexes. The pattern of fluorescence from GFP alone is identical in [PSI +] and [psi –] cells.

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from a Sup35-GFP fusion protein is diffusely distributed throughout the cytoplasm in [psi –] strains when expressed for a short time. Over the same time in [PSI +] cells, all detected fluorescence is concentrated in foci in the cytoplasm. This observation suggested that newly synthesized Sup35 protein is rapidly captured by the preexisting aggregates of Sup35 that are present in the [PSI +] cytoplasm, providing evidence for replication of the prion state. Indeed, epitope tagging of the endogenous Sup35 protein has recently confirmed that the newly made GFP-tagged Sup35 accumulates at endogenous, preexisting Sup35 foci (Liu and Lindquist, unpublished observation; Zhou et al., 2001). To test the hypothesis that Sup35-GFP coalescence was providing real-time visualization of the conversion process, the fusion protein was expressed in [psi –] cells for a longer period. Fluorescence begins to coalesce into discrete foci several hours after diffuse fluorescence is first detected. When these cultures are plated to media that selects for [PSI +] cells but not for the plasmid, [PSI +] cells can be isolated concomitant with the appearance of foci. Thus, the de novo appearance of these complexes of Sup35 is linked to the de novo acquisition of the [PSI +] state (Patino et al., 1996; Zhou et al., 2001).

IV. A MODEL FOR THE [PSI +] PHENOTYPE In the framework of the yeast prion hypothesis, the first molecular model of the [PSI +] phenotype was proposed (Fig. 3) (Patino et al., 1996; Paushkin et al., 1996). SUP35 is the yeast homolog of the eukaryotic release factor 3 (eRF3) and, together with SUP45 (eRF1), functions to terminate translation at nonsense codons (Stansfield et al., 1995; Zhouravleva et al., 1995). Reduction in Sup35 function caused by mutations in the SUP35 gene have the same phenotype as [PSI +], nonsense suppression (Chernoff et al., 1992b; Hawthorne and Leupold, 1974; Inge-Vechtomov and Andrianova, 1975). Unlike [PSI +], the phenotypes caused by mutations are inherited in a normal fashion. That is, they are recessive in crosses and segregate to only two of the four progeny of meiosis. However, the similarity in phenotype suggested that the [PSI +] phenotype might be due to a prion-like inactivation of Sup35: sequestration of Sup35 into large complexes in the [PSI +] cytoplasm would preclude its function in translation termination leading to an increase in stop codon read-through (nonsense suppression). The Sup35 protein complexes of [PSI +] cells are passed through the cytoplasm from mother cells to their daughters on cell division. These complexes

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FIG. 3. Model for the [PSI +] phenotype. The black line denotes a messenger RNA with a stop codon mutation in the coding sequence. In [psi –] cells (top), Sup35 (hatched spheres with black, crescent tails) is soluble and binds to Sup45 (black crescent). The Sup35/Sup45 complex promotes translation termination at all nonsense codons (stop). In [PSI +] cells (bottom), Sup35 forms aggregates in the cytoplasm through interaction of the N-terminal domain (black crescent). The level of soluble Sup35 available for binding to Sup45 is greatly reduced. Consequently, ribosomes (gray spheres) are able to read through nonsense codons at a low frequency (dashed arrow), leading to the production of full-length protein, which suppresses the effects of the mutation.

replicate by influencing newly synthesized protein to adopt the same state and join the complex. Thus, [PSI +] is inherited cytoplasmically. V. CRUCIAL RESIDUES IN THE REPLICATION OF PROTEIN STATES A. Sup35 Sequence Elements (N, M, C) Sup35 is a 685 residue protein that can be divided into three regions based on amino acid distribution and homology to other proteins (Fig. 4) (Kikuchi et al., 1988; Kushnirov et al., 1988; Wilson and Culbertson, 1988). The N-terminal region (N) is 124 residues and is very rich in glycine (G, 17%), tyrosine (Y, 16%), asparagine (N, 16%), and glutamine (Q, 28%). It also contains five imperfect repeats of the nonapep-

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FIG. 4. Schematic diagram of Sup35. Sup35 can be divided into three regions: N (aa 1–124), M (aa 125–254), and C (aa 255–685) (see text for details). Full-length protein can support viability, propagate [PSI +], and form amyloid fibers in vitro. The N region alone or in combination with M can propagate [PSI +] and form fibers; however, these regions cannot support viability in the absence of C.

tide QGGYQ(Q)QYNP. N is the prion-determining domain of Sup35. Amino acids 1–114 are necessary for the continued propagation of [PSI +] in vivo (Ter-Avanesyan et al., 1994). In addition, overexpression of N is sufficient to induce [PSI +] de novo in [psi –] strains (Derkatch et al., 1996; Ter-Avanesyan et al., 1993). Although crucial for [PSI +] inheritance, the N region is dispensable for normal growth (Kushnirov et al., 1990; Ter-Avanesyan et al., 1993). The M region (aa 125–254) is highly charged and also has an unusual distribution of residues. Glutamic acid (E, 18%) and lysine (K, 19%) are abundant, but there are no arginines, and aspartic acid (D, 5%) comprises only a minor fraction of the residues. In the context of an N deletion, M is dispensable for both viability and [PSI +] induction (Derkatch et al., 1996; Kushnirov et al., 1990; Ter-Avanesyan et al., 1993). The role of M in the maintenance of [PSI +] is unknown; however, the presence of M has a clear effect on the solubility of N. In vivo, a fragment containing N and M (NM) mirrors the behavior of fulllength Sup35; NM is soluble in [psi –] strains but insoluble in [PSI +] strains (Patino et al., 1996; Paushkin et al., 1996). In contrast, N alone

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is always insoluble in vivo and is incapable of switching between the states that define the [psi –] and [PSI +] phenotypes (Patino et al., 1996; Paushkin et al., 1996). The C region (aa 255–685) of Sup35 is responsible for the protein’s translation termination function. It is dispensable for both [PSI +] maintenance and induction but is essential for viability (Derkatch et al., 1996; Kushnirov et al., 1990; Ter-Avanesyan et al., 1994; Ter-Avanesyan et al., 1993). This region is evolutionarily conserved, is homologous to the yeast translational elongation factor EF–1α, contains several putative GTP binding sites, and complexes with Sup45 (Ebihara and Nakamura, 1999; Eurwilaichitr et al., 1999; Kikuchi et al., 1988; Kushnirov et al., 1988;Wilson and Culbertson, 1988). B. Two Regions in N Contribute to [PSI+] Inheritance A subfragment of the N region of Sup35 from aa 1–114 is both necessary and sufficient for the propagation and induction of [PSI +] in vivo (Derkatch et al., 1996; Kushnirov et al., 1990; Ter-Avanesyan et al., 1994; Ter-Avanesyan et al., 1993). A deletion within the N region removing aa 2–69 (∆BstEII) is unable to support [PSI+] propagation and cannot induce [PSI +] de novo in [psi –] strains when overexpressed, suggesting that specific sequences within N are crucial for [PSI ] metabolism in vivo (Derkatch et al., 1996; Ter-Avanesyan et al., 1993). More precise mutants isolated by several groups have begun to elucidate the specific characteristics of this region that are required for conversion between the [PSI +] and [psi –] states (Fig. 5). 1. Nonapeptide Repeats The first analyses of the SUP35 sequence made note of the unusual character of the N region, specifically the existence of five imperfect repeats of a glutamine-rich sequence (Fig. 5, see previously) (Kikuchi et al., 1988; Kushnirov et al., 1988; Wilson and Culbertson, 1988). Early work by Cox and Young had isolated a series of chromosomal mutations that lead to the irreversible loss of [PSI +]; these mutations were termed [PSI +]-no-more (Young and Cox, 1971). On further characterization, one of these mutations, PNM2, was genetically linked to the SUP35 gene and shown to be a glycine to glutamic acid change at residue 58 in the second nonapeptide repeat (Fig. 5) (Doel et al., 1994). Next, directed mutagenesis was used to analyze the importance of the repeats. Deletion of repeats number two through five (aa 57–93; ∆R2–5) similarly leads to a loss of [PSI +] (Liu and Lindquist, 1999).

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FIG.5. Crucial sequence elements for [PSI +] propagation. Shown is the amino acid sequence of the N region of Sup35. Boxed regions correspond to the five nonapeptide repeats implicated in [PSI +] inheritance (Derkatch et al., 1996; Liu and Lindquist, 1999; Ter-Avanesyan et al., 1993). Arrows indicate positions at which mutations were isolated that act as antisuppressors or [PSI +]-No-More mutations when expressed from a plasmid in cells with a wild-type copy of SUP35 in the genome (DePace et al., 1998). The underlined sequence can be replaced by polyglutamine and retain the capacity to co[???]

Moreover, expanding the second repeat (aa 57–65) to three copies (8 total repeats; R2E2) increases the spontaneous rate of [PSI +] appearance in [psi –] cells by more than three orders of magnitude (Liu and Lindquist, 1999). These observations link the integrity of the nonapeptide repeat region to the maintenance of [PSI+] in vivo and suggest that the ability to exist in the alternative [PSI +] and [psi –] states is intimately linked to the total number of these repeats. 2. Glutamines at the N-terminus A study by DePace et al. (1998) revealed a second area in the N region that was an important contributor to [PSI +] metabolism in vivo. Plasmids containing a SUP35 gene with a mutagenized N domain were tested for their ability to influence the [PSI +] phenotype by selecting for restoration of termination fidelity (or antisuppression). The mutations isolated clustered in a small region, aa 8–24 (Fig. 5). Some plasmid mutations masked the nonsense suppressor phenotype of the host strain without curing [PSI +] (antisuppressors; ASU); other mutations cured [PSI +] even though wild-type Sup35 was expressed in the same cells ([PSI +]-No-More; PNM). Notably, mutations in the nonapeptide repeats were not isolated in this screen, suggesting that the phenotypes of these ASU and PNM mutations may be more robust than the mutation isolated at residue 58 (PNM2). The vast majority of ASU and PNM mutations involved substitution of either glutamine or asparagine residues to charged amino acids such as serine and arginine (DePace et al., 1998). The polarity of the residues in this region, rather than the specific sequence, appears to be the most crucial aspect for [PSI ] metabolism; a variant of NM containing polyglutamine in place of aa 8–24 retains the ability to induce [PSI +] de novo in [psi –] cells when overexpressed (DePace et al., 1998).

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VI. MODELING [PSI +] IN VITRO A. Sup35 Self-Assembly In Vitro In vitro, Sup35 and fragments of the protein containing crucial regions self-assemble into large complexes. The process by which soluble protein is converted to this form has been linked to the mechanism of [PSI +] inheritance by several experiments and thereby provides an excellent model for studying the self-perpetuating change in state that the protein undergoes in vivo. Only fragments of Sup35 containing N undergo in vitro self-assembly, providing the first link between this process and the propagation of [PSI +] in vivo (Glover et al., 1997; King et al., 1997). Full-length Sup35 and the NM fragment exist in an unassembled form in physiological buffers for extended periods and eventually begin to slowly associate into large-molecular-weight complexes (Fig. 6) (Glover et al., 1997). Under the same conditions, M remains soluble for months. As was the case in the yeast cytoplasm, a fragment containing N alone is completely insoluble in physiological buffers in vitro but can form similar structures to those of NM and full-length Sup35p in the presence of denaturant (2 M urea or 40% acetonitrile) (Glover et al., 1997; King et al., 1997). Transmission electron microscopy of the complexes formed from these fragments revealed a highly ordered fibrillar structure (Fig. 6) (Glover et al., 1997; King et al., 1997). Fibers are stable structures and are resistant to solubilization with strong ionic detergent (2% w/v SDS) at room temperature (Serio et al., 2000). However, NM fibers can be disrupted with 2% (w/v) SDS at 100°C, indicating that covalent bonds do not contribute to this stability. The diameter of fibers is proportional to the size of the Sup35 fragment assembled: whole Sup35 (17 ± 2.0 nm), NM (11.5 ± 1.5 nm), and N (8.7 ± 0.9 nm) (Fig. 6) (Glover et al., 1997). If fibers of full-length Sup35 are incubated in high salt buffers, they adopt a more expanded structure revealing a central smooth rod with a diameter of 10.6 ± 1.0 nm surrounded by amorphous material along its length (Glover et al., 1997). In fibers, the N region is resistant to proteolysis, whereas the M region is accessible (Serio et al., 2000). Together, these observations are consistent with a defined topology in which the prion-determining N region forms the central core of the fiber and the M and C regions compose the exterior of the structure (Glover et al., 1997; Serio et al., 2000). Similar fibrillar structures have been observed for a number of proteins implicated in mammalian diseases including transthyretin, the Alzheimer’s disease-associated Aβ peptide, lysozyme, and the TSE-associ-

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FIG.6. Transmission electron microscopy of Sup35 fibers. Fibers of the NM fragment of Sup35 (A) or full-length Sup35 (B) were prepared and analyzed as described (Glover et al., 1997). Fibers prepared from Sup35 have a central axis, similar to that of fibers formed from the NM domain alone, with amorphous material, presumably the C domain, splayed out along the side. Scale bar, 100nm.

ated PrP, as well as for the yeast protein Ure2, associated with [URE3] (Serpell et al., 1997; Taylor et al., 1999). In the case of the mammalian proteins, fibers have been detected in vivo, and on the basis of this trait, the group is collectively known as protein amyloids. Amyloid fibers formed by members of this group share the ability to bind to the diagnostic dye Congo red and induce a shift in its absorption spectrum (Klunk et al., 1989). Fibers of both full-length Sup35 and the NM fragment bind to Congo red (4.4 binding sites per NM monomer, Kd = 250 nM) and induce the amyloid-characteristic spectral shift with a maximal difference at 540 nm (Glover et al., 1997). When viewed under polarized light, amyloid fibers bound to Congo red also exhibit green bi-refringence (Lansbury, 1992). Fibers formed from both N and NM also share this characteristic (Cashikar and Lindquist, unpublished observation, King et al., 1997). Members of the amyloid family have no sequence homology with each other, but fibers formed from proteins in this group share a similar secondary structure. Two characteristic reflections (at ~10 and 4.8 Å) are produced when fibers formed from any of the amyloid proteins are analyzed by x-ray diffraction (Sunde et al., 1997). This pattern defines the cross β-pleated sheet structure in which β sheets lie parallel to the long axis of the fiber, with the strands of each sheet oriented perpendicular to the fiber axis (Sunde et al., 1997). Fibers formed from the NM region of Sup35 share this structure (Serio et al., 2000); thus, Sup35 can be defined as an amyloidogenic protein.

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B. Replication of the Alternate Protein State in vitro The replication of conformational information in vivo has been demonstrated for Sup35 and strongly supports the hypothesis that [PSI +] is inherited through a protein-only mechanism (Patino et al., 1996). In vitro, the process of conformational replication has also been modeled for both the N and NM fragments of Sup35 (Glover et al., 1997; King et al., 1997). When diluted from denaturant, NM remains soluble, largely as a randomcoil, for several hours. After this lag phase, the protein is converted to amyloid fibers containing significant β-sheet structure as analyzed by circular dichroism (Glover et al., 1997). Conversion to the structured, assembled form is greatly accelerated by small quantities of preformed fibers, indicating that previously structured protein can influence the assembly of soluble protein (Glover et al., 1997; King et al., 1997). This in vitro seeded polymerization process has been linked to the mechanism of [PSI +] inheritance in vivo. Strikingly, the assembly of soluble NM in vitro can also be accelerated by the addition of lysates from [PSI +] but not from [psi –] cells (Glover et al., 1997). In a related series of experiments, [PSI +] lysates direct the in vitro self-assembly of fragments of Sup35 in [psi –] lysates, although the structure of these particles was not determined (Paushkin et al., 1997). These studies strongly link in vitro conversion to the in vivo propagation of [PSI +]. In several biological systems including the construction of viral capsids, bacterial flagella, and the amyloids associated with human disease, proteins are synthesized in a soluble state (S) that is distinct from the form that they adopt when assembled into a larger complex (A) (Caspar, 1980; Serpell et al., 1997). Three models have been proposed to explain the interplay between conformational conversion and assembly in these systems (Fig. 7). The models differ fundamentally in their predictions of the nature of the species that promotes conformational conversion (soluble monomers or solid-state protein), as well as the rate-limiting step (conformational change or polymerization). The first model, monomer-directed conversion (MDC) (Prusiner, 1982), predicts that A-state protein exists in solution but is rare. Interaction between soluble A- and S-state protein induces a conformational change, and two A-state monomers are released. The released A-state protein is then competent for self-assembly; that is, assembly is a consequence of the conversion process (the rate-limiting step). The second model, templated assembly (TA) (Griffith, 1967; Uratani et al., 1972), predicts that solid-state protein in the A-state catalyzes conformational conversion of S-state protein concomitant with assembly. Because S-state protein must convert conformationally to regenerate an assembly-competent surface, conversion is the rate-limiting step.

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FIG.7. Three models to explain the relationship between conformational change and assembly. S, state protein; light, jagged spheres; A, state protein: dark, smooth spheres; recently converted A-state protein: light, smooth spheres. See text for details of the models.

The third model, nucleated polymerization (NP) (Jarrett and Lansbury, 1993), predicts that the S- and A-states are in equilibrium in solution, but the S-state is predominant. Soluble A-state protein is stabilized by association with assembled A-state complexes. Thus, the rate-limiting step is formation of a polymerization-competent surface or nucleus rather than conformational conversion. In the case of NM, all available evidence suggests that fiber formation is nucleated by the solid-state and that fiber ends provide the crucial nucleating surface (Serio et al., 2000). Fibers direct the assembly of a vast excess of freshly added soluble NM within hours, and this nucleating activity is sedimentable at high speeds. Remarkably, when fibers are fractured by sonication, they have a greatly enhanced ability to nucleate conversion and assembly. Sonication does not alter the biochemical characteristics of fibers, nor does it increase the amount of soluble Astate protein; it simply fractures fibers into small pieces and thereby creates additional ends to serve as points for polymerization. Detailed time course analysis of NM fiber formation provided insight into the mechanism of nucleation (Serio et al., 2000). Specifically, a series of biochemical probes and microscopy techniques were used in concert to determine if preformed NM fibers acted as nuclei for con-

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formational change (TA) or polymerization (NP). The conformational change of soluble NM could not be separated from the assembly process in the presence of either intact or fractured fibers, and no pool of soluble but ordered (A-state) NM could be detected. These observations are consistent with the idea that preformed NM fibers act as nuclei for conformational change. Experiments in which the concentration of soluble protein is varied also suggest that NM assembly is limited by conformational change rather than polymerization. If fibers were assembled de novo via NP, the time for nucleus formation (lag phase) should vary inversely with the soluble monomer concentration, and the order of this reaction will be related to the number of A-state monomers that must associate to form a stable nucleus (Hofrichter et al., 1974; Jarrett and Lansbury, 1993). The lag phase for NM assembly in unnucleated reactions is independent of monomer concentration over a broad range (~0.1 to 50 µM) (Serio et al., 2000); thus, assembly of a nucleus from preexisting soluble A-state protein is not the rate-limiting step for NM assembly. However, if conformational conversion were the rate-limiting step for NM fiber formation, the time required for assembly onto a preformed nucleus would approach a constant time if the concentration of soluble protein exceeds that of polymerization surfaces (Caspar, 1980). This limit reflects the time required for S-state protein interacting with an A-state nucleus to conformationally convert and regenerate a polymerization competent surface. For NM, the length of the assembly phase is linearly dependent on monomer concentration below 5µM but plateaus above this point (Serio et al., 2000). Taken together these observations support the implications of the biochemical characterization of NM nucleated assembly discussed previously: NM conformationally converts to the A-state coincident with assembly, as suggested by the TA model. Although these analyses point to a mechanism of NM fiber growth in the presence of a preformed nucleus, they do not address the mechanism of de novo nucleus formation. Biochemical characterizations and concentration experiments suggest that A-state NM does not appreciably exist in solution, yet solutions of soluble NM protein will eventually form fibers (Glover et al., 1997). Unstructured oligomers of NM have been detected by multiple methods during the early stages of fiber formation (Glover et al., 1997; Serio et al., 2000). The high local concentration provided in these complexes may serve to stabilize protein that has converted to the A-state. Indeed, treatments that increase NM oligomerization in an unstructured state form fibers more rapidly, and conversely, decreasing NM oligomerization at early times in the assembly reaction results in a decreased rate of fiber formation (Serio et al., 2000).

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FIG.8. Nucleated conformational conversion model. S, state protein: jagged spheres; A-state protein: dark, smooth spheres; recently converted A-state protein: light, smooth spheres. See text for details of the models.

These observations suggest that NM nucleus formation occurs via a modified version of NP (Serio et al., 2000). The A-state is apparently so unstable or rare in solution that it cannot be detected directly by the biochemical means described previously or indirectly through a concentration-dependent lag phase. However, A-state protein may be stabilized in the context of an unstructured NM oligomer, or molten prenucleus. Once stabilized, this prenucleus converts to a bona fide Astate nucleus that is then competent to assemble and direct the conformational conversion of soluble S-state protein. Thus, the complete model that describes both NM nucleus formation, and its subsequent assembly onto preformed nuclei is a hybrid of both NP and TA. We refer to this model as nucleated conformational conversion or NCC (Fig. 8) (Serio et al., 2000). C. Biochemical Effects of N Mutations In vitro analysis of NM conversion reactions (discussed previously) suggests that conformational change occurs after unstructured protein interacts with a structured nucleus. It is currently unclear whether the protein-protein interaction domain in NM is overlapping or distinct

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from the region that adopts the β-sheet structure of mature fibers. Biochemical analyses of the collection of N mutants both in vivo and in vitro have begun to address the importance of both protein-protein interaction and β-sheet forming propensity in the switch between distinct protein states that forms the basis of [PSI +] inheritance. Sup35PNM2, Sup35∆R2–5, and many of the PNM and ASU mutations isolated by DePace et al. have a reduced capacity for association with wildtype Sup35 in vivo, and, conversely, Sup35R2E2 has an increased capacity for interaction with wild-type Sup35 (DePace et al., 1998; Kochneva-Pervakhova, 1998; Liu and Lindquist, 1999). The self-association properties of these mutants may form the basis of the molecular mechanism by which they alter [PSI] metabolism. For example, decreasing the efficiency of association between Sup35 molecules would eliminate or severely reduce replication of the prion state, leading to loss of the nonsense suppressor phenotype. Similarly, increasing the affinity of Sup35 molecules for each other could promote formation of the prion state much like the effect of excess wild-type Sup35 (Derkatch et al., 1996). This hypothesis is supported by in vitro studies with these proteins. The rate of self-assembly of Sup35PNM2, NM∆R2–5, and NMPNM or ASU is greatly reduced, whereas that of NMR2E2 is greatly accelerated compared with wild-type protein (DePace et al., 1998; Kochneva-Pervakhova, 1998; Liu and Lindquist, 1999). For both the collection of ASU and PNM mutations in NM and NMR2E2, the lag phases are decreased or increased, respectively, but assembly times are little affected (DePace et al., 1998; Liu and Lindquist, 1999). These observations suggest that protein-protein interaction is crucial for nucleus formation and that the N-terminal residues identified by mutational analyses play a crucial role in these interactions (DePace et al., 1998; Liu and Lindquist, 1999). Particular regions of N are also crucial for the protein-protein interactions that drive assembly onto a preformed nucleus both in vivo and in vitro. A series of related studies by three groups have demonstrated a species barrier for [PSI +] (Chernoff et al., 2000; Kushnirov et al., 2000; Santoso et al., 2000). The N regions of SUP35 cloned from several species of yeast can exist in different states (soluble vs. insoluble) both in vivo and in vitro, and remarkably, these proteins have also retained the capacity to promote their own assembly both in vivo and in vitro. However, the N region of Sup35 from one species cannot alter the assembly of the N region of Sup35 from another species even when present in the same cytoplasm or the same test tube. For Candida albicans and S. cerevisiae Sup35s, the crucial region for determining species specific protein-protein interactions localized to aa 8–26 (Santoso et al.,

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2000), residues previously identified by point mutagenesis as particularly sensitive to PNM and ASU mutations (DePace et al., 1998). Notably, a fusion protein linking GFP to a variant of S. cerevisiae NM containing a polyglutamine replacement of this region could be incorporated into aggregates present in the [PSI +] cytoplasm (DePace et al., 1998), suggesting that species differences in this region may actively disfavor the intrinsic propensity of polar residues to interact. In vitro studies of NM∆R2–5 and NMR2E2 fiber formation have also highlighted the importance of β-sheet forming propensity in self-assembly (Liu and Lindquist, 1999). R2E2 is the only known mutation in Sup35 that increases the spontaneous rate of [PSI +] appearance in [psi –] strains, and in vitro the same change results in an accelerated acquisition of β-sheet structure from the denatured state (Liu and Lindquist, 1999). Conversely, ∆R2–5 has a decreased propensity to adopt the β-sheet structure that is characteristic of NM fibers; the assembly time is greatly increased with this mutant, indicating that even in the presence of a structured nucleus conformational change is retarded (Liu and Lindquist, 1999). Although mutagenesis and domain swaps between homologs have highlighted the strong contributions of both interaction and conformation to both the formation and propagation of [PSI +] in vivo and amyloid in vitro, it is as yet unclear whether these attributes can be disentangled. These processes may result in an “induced fit” in which interaction promotes structure acquisition, which in turn stabilizes interaction. The propensity and stability of such assemblages have profound implications for continued propagation of [PSI +] and the ability of Sup35 to exist in multiple states and will undoubtedly be the focus of much future work. VII. REGULATION OF [PSI ] METABOLISM BY MOLECULAR CHAPERONES Although the tertiary structure of a protein is generally believed to be dictated by its primary sequence of amino acids (Anfinsen, 1967), some proteins are unique in that they are predicted to adopt at least two stable, structurally distinct states. Conditions that favor one state or the other have broad implications for the development of protein aggregation-associated diseases such as the prion diseases (Creutzfeld-Jakob, fatal familial insomnia, Gerstmann-Straussler-Scheinker), the systemic amyloidoses, Alzheimer’s disease, Parkinson’s disease, and Huntington’s disease (Serpell et al., 1997; Wickner et al., 1999). Mutations that destabilize the native state of such proteins (e.g., transthyretin and

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lysozyme) have been correlated with an increased incidence of disease (Lansbury, 1999). Molecular chaperones act posttranslationally to ensure structural quality control within the cell (Wickner et al., 1999). However, interactions between molecular chaperones and proteins capable of stably adopting alternate states may not always direct forward folding to the native or functional form (Lansbury, 1999). The disaggregating capacity of some members of this group, particularly the Clp/Hsp100 family, may promote acquisition of the alternate or disease-associated conformation by allowing the protein to reach a conformational transition state or facilitating the replication of this state once present. The functional interplay between this group and other chaperone systems such as the DnaK/Hsp70 and DnaJ/Hsp40 families is likely to further modulate the equilibrium between conformational isoforms in vivo and, thus, disease progression (Glover and Lindquist, 1998; Mogk et al., 1999; Zolkiewski, 1999). The available genetic and biochemical evidence indicates that [PSI +] arises from a self-perpetuating change in the physical state of Sup35 (see previously). In this system, molecular chaperones present in the yeast cytosol have been genetically implicated in transitions between Sup35 states, resulting in heritable changes in phenotype. HSP104 was isolated in a genetic screen for factors that increased the frequency of [PSI +] curing (Chernoff et al., 1995), and both SSA1 and SSB, members of the Hsp70 family, have subsequently been demonstrated to modulate the activity of HSP104 in [PSI] metabolism. Before discussing their roles in this capacity, we will briefly review their well-characterized roles in posttranslational quality control. A. HSP104 1. Role in Induced Stress Responses In yeast, tolerance to extreme environmental stresses can be induced by previous exposure to a more moderate stress (Parsell et al., 1993). For example, when yeast cultures are subjected to a sudden severe heat stress (25°C → 50°C), viability is rapidly lost within the first 5 minutes (Sanchez and Lindquist, 1990). However, when cultures are exposed to mildly elevated temperatures (37°C) before the severe stress, survival is greatly increased (Sanchez and Lindquist, 1990). This phenomenon is referred to as induced thermotolerance, and HSP104 is a crucial factor mediating the response (Sanchez and Lindquist, 1990). Although strains that are deficient in HSP104 grow normally at all temperatures, they have a 100- to 1000-fold reduced survival rate compared with wild-

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type strains when subjected to a severe thermal stress following a conditioning pretreatment (e.g., 25°C → 37°C → 50°C) (Sanchez and Lindquist, 1990). HSP104 is also required for tolerance to diverse forms of stress including ethanol and sodium arsenite, and in both cases, preexposure to low levels of these agents increases survival at higher levels (Piper, 1995; Sanchez et al., 1992). Moreover, exposure to one type of stress (e.g., ethanol) increases tolerance to other forms of stress (e.g., temperature) (Sanchez et al., 1992). Hsp104 is present at a very low level in log phase fermenting cultures grown at normal temperatures (Sanchez and Lindquist, 1990). Under these conditions, Hsp104 mutations have no noticeable effect on growth. Its concentration increases with exposure to stress (heat, ethanol), a switch to respiratory metabolism, entry into stationary phase growth, and sporulation (Sanchez and Lindquist, 1990; Sanchez et al., 1992). Compared with logarithmically growing cultures in glucose, cultures grown in nonfermentable carbon sources or those that have entered stationary phase exhibit a higher level of basal thermotolerance (e.g., without a conditioning pretreatment), providing another link between tolerance and Hsp104 (Sanchez et al., 1992). 2. Genetic Interaction with Hsp70 The S. cerevisiae genome encodes several members of the DnaK/Hsp70 family of molecular chaperones that have been grouped into four subfamilies SSA-SSD; the most heat-inducible members of this family are encoded by the SSA genes (Boorstein et al., 1994; Craig, 1990; Craig et al., 1995). Some members of the SSA subfamily are abundant at normal temperatures; others are strongly induced by stress. These proteins have overlapping functions that are essential for normal growth; individual members can be eliminated by mutation, but deletion of all Ssa proteins is lethal. Ssa1 promotes the refolding of chemically denatured substrates in vitro and acts with Ydj1, a member of the DnaJ/Hsp40 family, to prevent aggregation (Bush and Meyer, 1996; Cyr, 1995; Cyr and Douglas, 1994). In vivo, Hsp70 likely binds unfolded or partially unfolded proteins following environmental stress, preventing their aggregation. At normal temperatures, Ssa proteins bind nascent chains on polysomes and a variety of proteins in the process of transport and assembly (Craig et al., 1995). A series of in vivo and in vitro experiments (see later) have demonstrated a functional cooperation between Hsp104 and Ssa proteins. The role of Hsp104 in induced thermotolerance in yeast is most apparent at longer exposures to extreme heat stress (Sanchez and Lindquist, 1990). At shorter incubation times, wild-type and HSP104-deficient strains sur-

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vive equally well, suggesting that other factors contribute at least transiently to induced thermotolerance (Sanchez and Lindquist, 1990). Indeed, members of the Hsp70 family of proteins (Ssal, Ssa3, and Ssa4) in yeast play a role in induced thermotolerance under these conditions (Sanchez et al., 1993). The action of the Ssa proteins is not redundant with Hsp104, however. The Ssa proteins do not increase survival after longer thermal stresses even when over-expressed in hsp104 strains (Sanchez et al., 1993), and eliminating the stress-inducible members of the family has no effect on induced thermotolerance in the presence of HSP104 (Werner-Washburne et al., 1987). However, Hsp104 becomes crucial for growth in the absence of the constitutively expressed HSP70 genes (Sanchez et al., 1993). Thus, deficiencies in either Hsp70 or Hsp104 uncover temperature-dependent roles for the other factor. 3. Hsp104 Function Hsp104 is a highly conserved member of the Hsp100/Clp family of proteins (Parsell et al., 1991; Schirmer et al., 1996), but it has a unique role in protecting cells from different forms of stress. Deletion of HSP104 does not globally alter protein synthesis (Parsell et al., 1993), nor does it increase the rate of protein turnover (Parsell et al., 1994b; Parsell et al., 1993). Rather, Hsp104 functions in vivo to mediate the disaggregation of insoluble protein aggregates (Parsell et al., 1994b). After severe thermal stress, Hsp104 is required for both reactivation of a reporter enzyme and resolution of massive electron dense aggregates present in both the nucleus and cytoplasm of yeast cells (Parsell et al., 1994b). The function of Hsp104 in the rescue of heat-damaged proteins is dependent on two Walker-type nucleotide binding sites, NBD1 and NBD2 (Parsell et al., 1994b) and appears to be direct; Hsp104 is targeted to heat-induced cytoplasmic and nuclear aggregates (Kawai et al., 1999). NBD1 dominates the ATP hydrolytic activity of Hsp104 in vitro (Km ~ 5 mM, Vmax ~ 2 nmol min–1 µg–1) (Schirmer et al., 1998), whereas NBD2 mediates Hsp104 hexamerization (Parsell et al., 1994a). When analyzed by transmission electron microscopy, Hsp104 hexamers adopt a ringlike structure (diameter ~15.5 nm, central pore ~2.5 nm (Schirmer et al., 1996)) similar to those formed by other members of this family including ClpA (Kessel et al., 1995). A role for Hsp104 in the rescue of aggregated protein is unique among molecular chaperones, and this activity has been modeled in vitro (Glover and Lindquist, 1998). Together, purified Hsp70 (Ssa1) and Hsp40 (Ydj1) suppress the aggregation of chemically denatured reporter proteins but can only mediate the refolding of non-native

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monomeric protein. Larger complexes of aggregated, denatured protein cannot be salvaged by Hsp70 and Hsp40 in vitro. Hsp104 cannot prevent the aggregation of chemically denatured protein, but together with Hsp70 and Hsp40, Hsp104 rescues protein that has aggregated into larger complexes. As is the case in vivo, the in vitro activity of Hsp104 is dependent on the integrity of both NBD1 and NBD2, as well as the presence of ATP. Thus, Hsp104 mediates the reactivation of substrates that are otherwise refractory to the action of Ssa1/Ydj1 (Glover and Lindquist, 1998). Other members of the Hsp100/Clp family of proteins have subsequently been demonstrated to act in similar ways. The E. coli proteins ClpB (Hsp104 homolog), DnaK (Hsp70 homolog), and DnaJ (Hsp40 homolog) cooperate in vitro to reverse the aggregation of both chemically and thermally denatured proteins (Mogk et al., 1999; Zolkiewski, 1999). Another member of this family, ClpA, has recently been demonstrated to have unfoldase activity using a native monomeric protein as substrate (Weber-Ban et al., 1999). This activity, if shared by other members of the Hsp100/Clp family of proteins, may be crucial to resolve complexes of denatured, aggregated protein. 4. Role in [PSI +] Maintenance and Inheritance HSP104 was isolated in a genetic screen for factors that increased the frequency of [PSI +] curing (Chernoff et al., 1995). Extracopy HSP104 causes the reversible loss of [PSI +] in yeast strains (Chernoff et al., 1995), and conversion to the [psi –] state is accompanied by a change in Sup35 physical state from insoluble to soluble (Patino et al., 1996; Paushkin et al., 1996). Introduction of two nonsense codons at the beginning of the HSP104 gene blocks [PSI +] curing, indicating that Hsp104 protein is required for this effect (Chernoff et al., 1995). When placed under the regulation of an inducible promoter, HSP104 cures [PSI +] only in response to the induction stimulus (Chernoff et al., 1995). Notably, transient overexpression of Hsp104 is sufficient to cure [PSI +], and when Hsp104 levels are returned to normal, the [psi –] state persists (Chernoff et al., 1995). Thus, a transient increase in Hsp104 levels is sufficient to induce a heritable change in phenotype in yeast. When Hsp104 variants containing mutations in both of the nucleotide binding sites (e.g., NBD1 and NBD2) are overexpressed in [PSI +] strains that are also expressing wild-type Hsp104, [PSI +] is often cured (Chernoff et al., 1995). Notably, ablation of either site alone cures [PSI +] in some genetic backgrounds but not others, possibly implicating additional factors in the maintenance of [PSI +] (Chernoff

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et al., 1995; Newman et al., 1999; Patino et al., 1996). The observation that these mutated Hsp104s cured [PSI +] in the presence of wild-type Hsp104 suggested that inactivation of Hsp104, by the formation of mixed hexamers in this case, was responsible for [PSI +] loss. Indeed, Hsp104 disruption cures [PSI +] (Chernoff et al., 1995), and steady-state pools of Sup35 protein shift from the insoluble to the soluble state (Patino et al., 1996). Unlike strains cured of [PSI +] by extracopy. HSP104, [psi –] hsp104 strains cannot reacquire [PSI +] and are thus considered [PSI +]-no-more (Chernoff et al., 1995). Hsp104 also acts to modulate conversion between the soluble and insoluble states of other protein aggregate-associated disease factors, such as huntingtin and PrP (DebBurman et al., 1997; Krobitsch and Lindquist, 2000). When expressed in the yeast cytosol, huntingtin protein remains soluble, but when disease-associated variants of huntingtin containing expansions of polyglutamine (poly-Q) are expressed, they aggregate over the course of a few hours. The formation of these poly-Q dependent huntingtin aggregates requires the presence of Hsp104 (Krobitsch and Lindquist, 2000). In vitro, Hsp104 also increases the efficiency of conversion of PrP from a protease-sensitive form (PrPsen) to a protease-resistant form (PrPres). This effect is dependent on partially denatured infectious protein isolated from hamster brain (PrPSc) but does not require NBD1, NBD2, or ATP (DebBurman et al., 1997). These observations indicate that the Clp/Hsp100 family of chaperones has the capacity to influence diverse, unrelated proteins that are linked only by their capacity to exist in multiple, stable conformational states. Hsp104 likely mediates conversions between [psi –] and [PSI +] and PrPsen and PrPres by direct interaction. In the presence of Hsp104, Sup35 adopts an alternate fold by circular dichroism, and these solutions exhibit increased light scattering (Schirmer and Lindquist, 1997). The ATPase activity of Hsp104 is inhibited by both full-length Sup35 and the NM fragment in vitro (Schirmer and Lindquist, 1997). Hsp104 similarly alters the folding of PrP and Aβ, and these amyloidogenic substrates inhibit the ATPase activity of Hsp104 (Schirmer and Lindquist, 1997). Taken together, these observations link a change in substrate conformation to a change in Hsp104 ATPase activity. 5. Models for Hsp104 Regulation of [PSI] Metabolism Two models have been proposed to explain the genetic interaction between Hsp104 and [PSI +] in vivo (Fig. 9) (Kushnirov and TerAvanesyan, 1998; Lindquist and Schirmer, 1999; Patino et al., 1996; Paushkin et al., 1996). The models differ in the crucial role Hsp104 plays in the maintenance of [PSI +]. The first model predicts that Hsp104 is

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FIG.9. Models of Hsp104 function in [PSI] maintenance and curing. Sup35, striped spheres with black, crescent tails; Hsp104, gray ovals; Sup35 transition state, Sup35*; Model 1 as per Patino et al. (1996); model 2 as per Kushnirov and Ter-Avanesyan (1998). See text for details of the models.

required for nascent or preexisting Sup35[psi –] to reach a transition state (conformational or oliogomeric) that is required for conversion to Sup35[PSI+] (Lindquist and Schirmer, 1999; Patino et al., 1996). The second model suggests that Hsp104 partially disaggregates existing Sup35[PSI+] complexes, thereby increasing the number of complexes within the cytoplasm while reducing their average size (Kushnirov and Ter-Avanesyan, 1998; Paushkin et al., 1996). According to the second model, smaller complexes would more efficiently promote conversion of newly synthesized Sup35 and would be more efficiently transmitted to daughters on cell division. Unfortunately, little information is available to distinguish between these models, but in potential support of the first, fluorescence from Sup35-GFP fusion proteins remains diffusely distributed throughout hsp104 cells even when overexpressed to extremely high levels, indicating that Hsp104 is required for Sup35 to aggregate in the first place (Liu and Lindquist, unpublished observation). With regard to [PSI +] curing by excess Hsp104, the models differ as well. Both models suggest that Hsp104 might disaggregate Sup35[PSI+] complexes at a rate that eventually out-competes conversion of newly synthesized Sup35 (Kushnirov and Ter-Avanesyan 1998; Lindquist and

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Schirmer, 1999; Patino et al., 1996; Paushkin et al., 1996). The first model also suggests another possibility (Lindquist and Schirmer, 1999; Patino et al., 1996). A balance between Hsp104 and Sup35 levels appears to be crucial in maintaining a stable [PSI +] state; too much or too little Hsp104 leads to curing. If oligomeric complexes are an obligate intermediate in the addition of new Sup35 protein to preexisting [PSI +] aggregates, then the formation of such complexes might require an intermediate concentration of Hsp104. That is, Hsp104 would be required to produce the crucial conformational transition-state of Sup35, but this state might be unstable in the absence of interaction with other proteins in the same state. When Hsp104 concentrations are limiting, the high local concentration of converted Sup35 substrates in the vicinity of an individual Hsp104 protein would then be crucial for oligomer formation. This model is supported by in vitro results suggesting oligomeric intermediates are involved in conformational conversion (see previously). Again, little information is currently available to distinguish between these models. One difficulty is that [PSI +] is normally detected by colony formation on selective media, which temporally distances the assay from the experimental manipulations that affect it. Testable differences between the two models do exist, however. According to the second model, excess Hsp104 would block replication of Sup35[PSI+] without affecting preexisting Sup35 aggregates. [PSI +] would eventually be lost from dividing cultures as preexisting [PSI +] complexes are diluted out. Thus, future studies monitoring the fate of preexisting Sup35 and nascent Sup35 in cells overexpressing Hsp104 may resolve this issue. B. SSA1 Given the functional interactions between Hsp104 and Ssal both in vivo and in vitro with denatured substrate proteins (Glover and Lindquist, 1998; Sanchez et al., 1993), it seemed likely that Ssal might play a role in [PSI] metabolism as well. Ssal is a member of the yeast Hsp70 family that is constitutively expressed at normal temperatures and is induced 2- to 3-fold after heat stress (Werner-Washburne and Craig, 1989; Werner-Washburne et al., 1987). Ssal expression is not always coordinately regulated with that of Hsp104 (Sanchez and Lindquist, 1990; Werner-Washburne et al., 1989, Werner-Washburne and Craig, 1989; Werner-Washburne et al., 1987), but overexpression of Ssal does alter [PSI] metabolism (Chernoff et al., 1995; Newman et al., 1999). Extra-copy SSA1 plasmids interfere with the ability of extra-copy HSP104 plasmids to cure [PSI +] (Newman et al., 1999). [PSI +] cells over-

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expressing Hsp104 alone have an increased amount of soluble Sup35, but elevated levels of Ssal counteract this effect (Newman et al., 1999). Moreover, excess Ssal increases the strength of the [PSI +] phenotype, an observation consistent with a decreased level of soluble Sup35 (Newman et al., 1999). These observations, taken together, suggest that Ssal counteracts the role of Hsp104 in [PSI +] curing. It is unclear, however, whether Hsp70 antagonism is direct or indirect. Modulation of Hsp70 levels have profound effects on the expression of many other proteins in the cell (Werner-Washburne et al., 1987). Moreover, the Hsp70 genes are essential for viability; thus, [PSI ] metabolism cannot be assessed in Hsp70-deficient strains (Werner-Washburne et al., 1987). Although the mechanism of action is unclear, the antagonism between Hsp104 and Hsp70 may have profound importance for our understanding of [PSI +] maintenance and inheritance. [PSI +] is stable to both heat shock and meiotic division, two physiological states that are accompanied by a massive induction of Hsp104 (Sanchez and Lindquist, 1990; Sanchez et al., 1992, Singh, 1979; Tuite et al., 1981). If elevated Hsp104 levels are sufficient to cure [PSI +] under normal growth conditions, why is [PSI +] protected during these stresses? One possible explanation is that, in addition to Hsp104, the levels of other chaperones are also increased by these treatments. These factors may protect [PSI +] by diverting Hsp104 to other targets and/or directly influencing the physical state of Sup35. When Hsp104 levels are increased in isolation of these other factors, the delicate balance between Sup35[PSI+] and Sup35[psi –] may be more susceptible to change. For example, the ability of Hsp70 to bind to unfolded proteins might stabilize conformational transition-state intermediates, reducing the diluting effect of Hsp104 overexpression. Another possibility is that both heat stress and sporulation are accompanied by a cessation of cellular division. If excess Hsp104 simply acts to block replication of Sup35[PSI+], several divisions would be required to dilute preexisting Sup35[PSI+] complexes. In the absence of division, these complexes would remain in the cytoplasm unperturbed until growth conditions returned to normal. At this point, Sup35[PSI+] replication could resume without an apparent change in [PSI] status. C. SSB The SSB subfamily of Hsp70 in S. cerevisiae has also recently been implicated in [PSI] metabolism (Chernoff et al., 1999). This group consists of two members, Ssb1 and Ssb2, which have identical amino acid sequences except at four positions (Boorstein et al., 1994). SSB proteins are not

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essential, but strains that are completely deficient (ssb1 ssb2 deletions) are sensitive to protein synthesis inhibitors. The Ssbs have been demonstrated to associate with both the ribosome and the nascent polypeptide chain (Nelson et al., 1992; Pfund et al., 1998). Together these observations suggest a role for the Ssbs in cotranslational protein folding. In contrast to the antagonistic interplay between excess Ssal and Hsp104, excess Ssb protein enhances the efficiency with which excess Hsp104 cures cells of [PSI +] (Chernoff et al., 1999). In addition, the rate of [PSI +] de novo induction by extra copy SUP35 is enhanced in ssb1 ssb2 strains, and [PSI +] mediated nonsense suppression is more efficient in ssb1 ssb2 strains in comparison with wild-type (Chernoff et al., 1999). These results suggest that Hsp104 and the Ssbs cooperate to tip the balance in favor of the [PSI +] conformation of Sup35 and intriguingly highlight the importance of cotranslational events in the conversion process. The mechanism by which the Ssbs act in concert with Hsp104 is unclear. Like Hsp104, the Ssbs could either act to block formation of a critical folding or oligomeric intermediate necessary for conversion, or, alternatively, they could act in concert to refold Sup35 that has been liberated from [PSI +] complexes present in the cytosol. Intriguingly, Ssb1 has been linked to efficient protein turnover in yeast, providing another potential but as yet unexplored pathway for the regulation of [PSI] metabolism in vivo (Ohba, 1994; Ohba, 1997). VIII. SUMMARY Biochemical characterization of the yeast prions has revealed many similarities with the mammalian amyloidogenic proteins. The ease of generating in vivo mutations in yeast and the developing in vitro models for [PSI +] and [URE3] circumvent many of the difficulties of studying the proteins linked to the mammalian amyloidoses. Future work especially aimed at understanding the molecular role of chaperone proteins in regulating conversion as well as the early steps in de novo formation of the prion state in yeast will likely provide invaluable lessons that may be more broadly applicable to related processes in higher eukaryotes. It is important to remember, however, that there are clear distinctions between disease states associated with amyloidogenesis and the epigenetic modulation of protein function by self-perpetuating conformational conversions. Amyloid formation is detrimental to mammals and is likely selected against, providing a possible explanation for the late onset of these disorders (Lansbury, 1999). In contrast, the known yeast prions are compatible with

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normal growth and, if beneficial to the organism, may be subject to evolutionary pressures that ultimately maximize transmission. In the prion proteins examined to date, distinct domains are responsible for normal function and for the conformational switches producing a prion conversion of that function. Recent work has demonstrated that the prion domains are both modular and transferable to other proteins on which they can confer a heritable epigenetic alteration of function (Edskes et al., 1999; Li and Lindquist, 2000; Patino et al., 1996; Santoso et al., 2000; Sondheimer and Lindquist, 2000). That is, prion domains need not coevolve with particular functional domains but might be moved from one protein to another during evolution. Such processes may be widely used in biology. Mechanistic studies of [PSI +] and [URE3] replication are sure to lay a foundation of knowledge for understanding a host of nonconventional genetic elements that currently remain elusive. REFERENCES Anfinsen, C.B. 1967. Harvey Lect 61, 95–116. Boorstein, W.R., Ziegelhoffer, T., and Craig, E.A. 1994. J. Mol. Evol. 38, 1–17. Bush, G.L., and Meyer, D.I. 1996. J. Cell. Biol. 135, 1229–1237. Caspar, D.L. 1980. Biophys. J. 32, 103–138. Chernoff, Y., Galkin, A., Lewitin, E., Chernova, T., Newnam, G., and Belenkiy, S. 2000. Mol. Microbiol. 35, 865–876. Chernoff, Y.O., Derkach, I.L., and Inge-Vechtomov, S.G. 1993. Curr. Genet. 24, 268–270. Chernoff, Y.O., Inge-Vechtomov, S.G., Derkach, I.L., Ptyushkina, M.V., Tarunina, O.V., Dagkesamanskaya, A.R., and Ter-Avanesyan, M.D. 1992a. Yeast 8, 489–499. Chernoff, Y.O., Lindquist, S.L., Ono, B., Inge-Vechtomov, S.G., and Liebman, S.W. 1995. Science 268, 880–884. Chernoff, Y.O., Newnam, G.P., Kumar, J., Allen, K., and Zink, A.D. 1999. Mol. Cell. Biol. 19, 8103–8112. Chernoff, Y.O., Ptyushkina, M.V., Samsonova, M.G., Sizonencko, G.I., Pavlov, Y.I., TerAvanesyan, M.D., and Inge-Vechtomov, S.G. 1992b. Biochimie 74, 455–461. Cox, B. 1965. Heredity 20, 505–521. Cox, B.S., Tuite, M.F., and McLaughlin, C.S. 1988. Yeast 4, 159–178. Cox, B.S., Tuite, M.F., and Mundy, C.J. 1980. Genetics 95, 589–609. Craig, E. 1990. Regulation and function of the HSP70 multigene family of Saccharomyces cerevisiae. In “Stress proteins in Biology and Medicine,” (R. Morimoto, A. Tissieres, and C. Georgopoulos, Eds.), pp. 301–321. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Craig, E., Ziegelhoffer, T., Nelson, J., Laloraya, S., and Halladay, J. 1995. Cold Spring Harbor Symp Quant Biol. 60, 441–449. Cyr, D.M. 1995. FEBS Lett. 359, 129–132. Cyr, D.M., and Douglas, M.G. 1994. J. Biol. Chem. 269, 9798–9804. DebBurman, S.K., Raymond, G.J., Caughey, B., and Lindquist, S. 1997. Proc. Natl. Acad. Sci. U.S.A. 94, 13938–13943.

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AUTHOR INDEX

A

Anderson, R. G. W., 201, 211, 224, 225, 228 Anderson, R. M., 1, 21 Andrianova, V., 341, 362 Anfinsen, C. B., 352, 361 Ang, L. C., 200 Angeretti, N., 23, 197, 200, 306 Anil-Kumar, 62, 79 Antoniou, M., 166 Appel, T. R., 51, 54, 101, 102, 162, 164, 167, 269 Arbustini, E., 268 Arcamone, F., 263 Ardini, E., 238, 239, 240, 241, 263, 264 Arnold, J. E., 101, 102, 216, 224 Arnsdorf, M. F., 363 Arunachalam, B., 95, 102 Asakura, S., 364 Asante, E., 22, 80, 154, 156, 161, 299, 303, 305 Asante, E. A., 273–304 Ascari, E., 268 Asher, D. M., 304 Askanas, V., 235, 263 Atarashi, R., 227, 308 Atzori, M., 199 Aucouturier, P., 169, 200, 270 Aufderheide, K. J., 327, 331 Austin, A. R., 21 Autenried, M., 80 Autenried, P., 22, 103, 264, 305, 330 Auth, D., 238, 239, 263 Autilio, G. L., 264 Autilio-Gambetti, L., 22, 103, 167, 225, 308 Avila, J., 199 Avoni, P., 308 Avraham, D., 26, 201, 228 Avraham, D. C. T. A. D., 270 Avrahami, D., 201, 228, 270 Awan, T., 200

Aagaard, C., 169, 201, 310 Aberth, W., 29, 52 Abildgaard, H. J. D., 82 Abraham, D. J., 362 Achermann, P., 81, 228, 271 Achermann, P. A., 311 Adams, J. M., 253, 263 Adjou, K., 165, 265 Adjou, K. T., 163, 164, 257, 260, 263, 265 Aebersold, R., 30, 52, 81, 167 Aebi, M., 80, 104, 331, 362 Aguet, M., 22, 80, 103, 197, 225, 264, 305, 330 Aguzzi, A., 21, 22, 23, 25, 80, 103, 105, 197, 200, 204, 228, 264, 270, 304, 305, 306, 309, 310, 330 Ahlstrom, K. R., 168 Aigle, M., 319, 329, 330 Aiken, J., 167, 268 Aiken, J. M., 79, 164 Aitchison, L., 267, 307 Akowitz, A., 162, 164 Albelda, S. M., 237, 263 Albrechtsen, R., 271 Algeri, M., 200 Allen, J. M., 304 Allen, K., 361 Alonso, D., 23, 116, 134, 135 Alonso, D. O. V., 107–134, 135, 136 Alper, T., 3, 21, 55, 56, 74, 79, 83, 102, 327, 330 Alperovitch, A., 27, 201, 311 Altman, R., 136 Altman, R. A., 268 Amann, E., 269 Amboldi, N., 268 Amico, C., 197 Anderson, R. G., 232, 241, 263, 271 367

368

AUTHOR INDEX

B Bacote, A., 304 Baest, J., 200 Bahr, U., 29, 52 Bai, Y. W., 64, 79 Baker, C. A., 196 Baker, H. F., 307, 308 Baldwin, M., 25, 80, 81, 103, 104, 105, 108, 109, 135, 136, 167, 198 Baldwin, M. A., 21, 23, 24, 29–52, 30, 33, 34, 36, 39, 41, 42, 43, 45, 47, 50, 52, 53, 54, 80, 82, 84, 102, 103, 105, 135, 136, 137, 140, 141, 166, 167, 168, 169, 198, 199, 201, 227, 263, 307, 309 Ball, H., 82, 166 Ball, H. L., 24, 50, 54, 105, 198 Bamborough, P., 23, 135, 272 Bandiera, T., 169, 270 Banfield, B. A., 26 Bao, Q., 237, 263 Barbanti, P., 100, 102 Barber, M., 29, 52 Barbieri, B., 261, 263 Barenholz, Y., 227 Barker, K. T., 264 Baron, G., 139–164 Barondes, S. H., 266 Barra, D., 105 Barron, R., 25, 308 Barrow, C. J., 135, 198 Barry, R. A., 23, 30, 52, 81, 167, 225, 306 Barsky, S. H., 269 Bartels, C., 62, 79 Bartlam, M., 364 Bartz, J., 167 Bartz, J. C., 77, 79, 157, 161, 164 Baruzzi, Q., 308 Basler, K., 30, 53, 57, 79, 108, 135, 274, 302, 304 Bass, N. H., 185, 196 Bastidas, R. B., 82, 136 Bax, A., 61, 62, 69, 72, 79, 81 Baxter, H. C., 23, 198 Baxter, N. J., 80, 103, 136 Baybutt, H., 25, 308 Bazan, J. F., 57, 79 Beck, K., 237, 239, 254, 263 Becker, J., 364 Beckstead, J. H., 225

Begueret, J., 325, 330, 332 Behringer, R. R., 304 Beisson, J., 327, 330 Beisson-Schecroun, J., 326, 330 Bekker, T., 80, 198 Belenkiy, S., 361 Bellini, O., 263 Bellotti, V., 103, 265, 268 Belotti, D., 266 Belt, P. B. G. M., 15, 16, 21, 165 Ben, S. S., 265 Bendheim, P. E., 6, 21, 139, 146, 161, 165, 168, 200, 204, 224, 249, 263, 304, 306 Bennett, A. D., 263 Bensasson, S. A., 23 Beringue, V., 165, 167, 263, 265, 267, 307 Berkenbosch, F., 201 Bernhard, 304 Berti, A., 266 Bertram, J. F., 267 Besinger, A., 10, 22, 57, 79, 205, 224 Bessen, R. A., 8, 11, 12, 21, 22, 25, 56, 79, 93, 102, 125, 135, 142, 145, 146, 151, 152, 153, 160, 161, 164, 165, 173, 197, 200, 215, 224, 328, 330 Betsholtz, C., 271 Beyreuther, K., 23, 30, 53, 103, 135, 166, 198, 268 Bhat, K. S., 22, 135, 165, 197, 305 Bigam, C. G., 82 Bignami, 185 Bilak, M., 263 Billeter, d. V. T., 265 Billeter, M., 26, 27, 62, 70, 79, 80, 81, 82, 85, 102, 104, 105, 123, 135, 136, 137, 168, 198, 200, 201, 262, 309 Billette de Villemeur, T., 225 Bird, T., 136, 199 Bird, T. D., 200, 201 Birkett, C., 22, 165, 197, 305 Birkett, C. R., 26, 166, 168, 169, 310 Biwersi, J., 263 Black, C., 226, 308 Blackburn, J. M., 332 Blaettler, T., 270 Blake, C. C., 103, 363, 364 Blattler, T., 3, 21, 310 Blättler, T., 279, 304 Blochberger, T. C., 8, 9, 21, 238, 263 Blockberger, T. C., 24, 166

369

AUTHOR INDEX

Bloomberg, G. B., 26 Bluethmann, H., 80, 264, 305, 330 Bockman, J. M., 225 Bohm, M., 199 Bolard, J., 260, 263, 268 Bolton, D. C., 4, 6, 21, 31, 53, 139, 146, 161, 165, 168, 200, 224, 249, 263, 304, 307 Boorstein, W. R., 354, 360, 361 Booth, D. R., 90, 103 Borchelt, D. R., 4, 5, 6, 21, 26, 30, 53, 81, 101, 103, 109, 135, 141, 165, 208, 209, 219, 220, 224, 225, 228 Bordoli, R. S., 29, 52 Bordoni, T., 263 Borkovich, K. A., 363 Bosque, P., 169, 201, 310 Bossers, A., 15, 21, 25, 104, 151, 152, 156, 157, 165, 168 Bostock, C., 24, 25, 308 Bostock, C. J., 22, 165, 166, 197, 263, 305 Boughey, A. M., 305 Bowman, K. A., 31, 53, 168, 200 Bradley, R., 1, 21, 22, 27, 80, 305 Brady, K., 226 Braginski, J. E., 264 Brajtburg, J., 268 Brandner, S., 3, 21, 23, 25, 103, 105, 197, 200, 270, 279, 300, 301, 303, 304, 306, 309, 310 Brasseur, R., 187, 197, 198, 200 Braun, U., 80, 136, 198 Braun, W., 79, 82 Brawerman, G., 238, 239, 263 Brentani, R. R., 266, 268 Brignoni, M., 270 Brimacombe, D. B., 231, 232, 255, 258, 263 Brinkley, B. R., 362 Brinster, R. L., 302, 304 Broach, J. R., 330 Brose, N., 226 Brown, C. R., 248, 263, 271 Brown, D. R., 10, 22, 23, 57, 74, 79, 102, 103, 178, 179, 189, 190, 192, 195, 197, 205, 210, 224, 225, 230, 232, 263, 264, 281, 300, 304 Brown, H. R., 224 Brown, K., 22, 264 Brown, P., 22, 23, 25, 55, 79, 103, 167, 198, 199, 200, 227, 288, 304, 306, 308 Bruce, M., 24, 265, 294, 297, 305

Bruce, M. E., 2, 10, 11, 22, 24, 156, 159, 165, 166, 173, 188, 197, 226, 293, 294, 297, 298, 299, 304, 305 Brunori, M., 105 Bruschi, M., 200 Bryant, P. K. III, 168 Buchmeier, M. J., 22, 165, 225 Buck, C. A., 237, 263 Buckle, B. M., 82 Budka, H., 199, 227 Bueler, H., 3, 17, 22, 229, 232, 264, 328, 330 Büeler, H., 56, 57, 80, 83, 96, 103, 105, 189, 197, 204, 225, 275, 278, 301, 302, 305, 309 Büeler, H. R., 306 Bugiani, M., 169, 270 Bugiani, O., 23, 25, 26, 105, 166, 169, 171–196, 197, 198, 199, 200, 201, 227, 228, 270, 306 Bukau, B., 363 Buller, R., 22, 225 Burkle, A., 23, 166, 198 Burlingame, A., 225 Burlingame, A. L., 29, 30, 33, 34, 36, 39, 41, 42, 43, 45, 50, 52, 53, 54, 105, 137, 227, 228 Burman, 153 Burton, D., 309 Burton, D. R., 82, 136 Buschmann, A., 293, 305 Bush, G. L., 354, 361 Butler, D. A., 212, 225, 307 Butler, P. J. G., 332 Buto, S., 238, 240, 264

C Cabral, A. L., 266 Cabry, J., 196 Cairns, D., 166 Calayag, M. C., 225 Callahan, M., 139–164 Calzolai, L., 27, 68, 75, 80, 82, 105, 137, 201 Camerino, A. P., 23, 135, 271, 306 Canciani, B., 169, 270 Candy, J. M., 24, 50, 53, 198, 226, 307 Canfield, S. M., 240, 264 Cantoni, L., 200 Cantù, L., 197, 200 Capellari, S., 22, 25, 103, 167, 199, 207, 225, 227, 309

370

AUTHOR INDEX

Cappai, R., 128, 135, 198 Cappellari, S., 199 Cardone, F., 102, 105 Carlin, B., 264 Carlson, G., 26, 81 Carlson, G. A., 25, 27, 168, 226, 268, 272, 296, 305, 308, 309, 310, 311, 332 Carmi, P., 268 Caron, M G., 269 Carp, R. I., 24, 166, 167, 169, 198, 200, 270, 297, 305, 306 Carr, S. A., 30, 52 Casaccia, P., 167, 267 Casanova, M., 267 Casazza, A. M., 263 Cashikar, A. G., 347, 363 Cashman, N., 236, 242, 243, 264 Cashman, N. R., 204, 224, 225 Caspar, D. L., 348, 349, 361 Caspi, S., 257, 258, 264 Cass, C., 136 Cassutti, P., 169, 270 Castellani, R., 25, 199, 309 Castle, B. E., 199 Castronovo, V., 237, 238, 239, 240, 263, 264, 265, 266, 269 Caughey, B., 3, 4, 5, 6, 8, 9, 17, 21, 22, 23, 24, 25, 50, 53, 57, 79, 80, 83, 84, 93, 102, 103, 104, 108, 109, 110, 123, 124, 130, 135, 136, 139–164, 165, 166, 167, 168, 197, 204, 206, 208, 209, 216, 219, 220, 222, 225, 227, 231, 255, 257, 258, 264, 265, 269, 313, 315, 330, 331, 362 Caughey, B. W., 22, 135, 141, 165, 171, 197, 274, 305 Caughey, W. S., 22, 135, 142, 146, 165, 168, 197, 257, 261, 264, 269, 305 Cayetano, C. J., 311 Cayetano, J., 23, 135, 271, 306 Cayetano-Canlas, J., 332 Cereghetti, G., 86, 91, 99, 103, 105 Ceroni, M., 26, 168, 227 Cervenakova, L., 23, 306 Cervini, M. A., 169, 270 Chabry, J., 9, 22, 24, 129, 135, 145, 152, 155, 156, 163, 165, 166, 197 Chait, B. T., 29, 47, 49, 52, 53, 225 Chang, W.-J., 211, 225 Chechenova, M. B., 362 Cheisa, R., 305 Chen, H. S., 272

Chen, H. Y., 308 Chen, L. B., 272 Chen, S. G., 12, 17, 18, 20, 22, 25, 100, 101, 103, 104, 105, 167, 168, 199, 208, 222, 225, 227, 231, 264, 308, 309 Cheng, N., 105, 332, 364 Cherifi, K., 165, 265 Chernoff, Y., 351, 361 Chernoff, Y. O., 245, 264, 319, 330, 336, 339, 341, 353, 356, 358, 359, 360, 361, 362, 363, 364 Chernova, T., 361 Chesebro, B., 3, 9, 15, 16, 19, 22, 24, 25, 108, 135, 136, 140, 141, 149, 159, 165, 167, 168, 197, 200, 213, 225, 227, 250, 269, 309, 331 Chiarle, R., 199 Chien, P., 104, 363 Chiesa, R., 23, 194, 197, 200, 213, 225, 286, 303, 306 Chishti, M. A., 226, 268 Chishti, Y. L., 308 Choi, E., 198 Choi, J. H., 198 Chong, A., 23, 26, 39, 53, 166, 169, 198, 228, 310 Chou, P. Y., 111, 116, 135 Chou, S. M., 185, 197 Chree, A., 22, 165, 197, 305 Christen, P., 50, 53, 198, 267, 307 Chung, A. E., 254, 264 Cibati, M., 167 Cioce, V., 264 Clarke, A. R., 24, 25, 26, 80, 103, 105, 136, 166, 169, 201, 225, 226, 264, 267, 305, 307, 308 Clarke, M. C., 21, 79, 102, 330 Clausse, N., 240, 264, 266 Clave, C., 330 Claysmith, A. P., 264, 269, 271 Clive, C., 22, 264 Cochran, E. J., 22, 103 Cochran, S. P., 31, 53 Cohen, F., 134, 135, 198, 308 Cohen, F. E., 23, 24, 25, 26, 27, 50, 52, 54, 80, 81, 82, 96, 102, 103, 104, 105, 111, 135, 136, 137, 166, 167, 168, 169, 197, 198, 199, 200, 201, 226, 228, 265, 266, 268, 269, 271, 272, 306, 307, 308, 309, 310 Colaco, C., 199

AUTHOR INDEX

Colello, R. J., 226, 308 Coleman, S., 305 Coligan, J. E., 271 Colling, S. B., 205, 225, 275, 305 Collinge, J., 2, 11, 13, 22, 24, 25, 26, 57, 77, 80, 81, 84, 93, 103, 105, 136, 146, 154, 156, 160, 161, 165, 166, 167, 169, 201, 205, 225, 228, 229, 264, 273–304, 305, 306, 307, 308, 309, 311 Collins, S. J., 135, 198 Colnaghi, M. I., 263, 264 Colominas, C., 39, 53, 81, 137, 168 Come, J. H., 24, 136, 167, 331 Conde, J., 336, 362 Cook, R. F., 226 Cooper, C., 21, 263 Cooper, C. M., 80, 104, 136, 166, 266 Cooper, T. G., 316, 317, 330, 332 Corrales, F. J., 82 Cortelli, P., 27, 137, 167, 169, 308, 309, 310 Cory, S., 253, 263 Coschigano, P. W., 317, 319, 330 Costa, M. D., 310 Cotman, C. W., 200 Cotter, R., 167 Coufal, F., 26, 81, 310 Courchesne, W. E., 317, 318, 330 Cousens, S., 22, 165, 197, 305 Cousens, S. N., 27, 201, 311 Coustou, V., 326, 330 Cox, B., 335, 336, 338, 344, 361, 364 Cox, B. S., 314, 318, 319, 330, 331, 332, 336, 361, 362, 363, 364 Cozzio, A., 270, 276, 278, 306, 310 Craig, E., 354, 361 Craig, E. A., 358, 361, 363, 364 Cramp, W. A., 21, 79, 102, 330 Craven, C. J., 24, 80, 103, 136, 166 Cresswell, P., 102 Crow, T. J., 305, 307, 308 Culberston, M. R., 342, 344, 364 Currie, J. R., 304 Curtis, D., 195, 197 Cyr, D. M., 354, 362

D D’ursi, P., 171–196 Da Costa, M., 26, 81, 105 Da, C. M., 272 Daggett, V., 23, 134, 135, 136, 306

371

Dagkesamanskaya, A. R., 310, 332, 361, 362, 363, 364 Dall’Ara, P., 169, 270 Damm, J. C., 263 Daude, N., 19, 22, 103, 213, 219, 220, 221, 225, 226 Davidson, M., 270 Davies, E., 265 Davis, S. C., 239, 240, 265 Dawson, M., 23, 27 De Carli, C., 200 De Fea, K. A., 23, 80, 103, 198, 218, 225, 226, 306 De Gioia, L., 181, 197, 200 de Vries, R., 21, 165 de, S. S., 268 DeArmond, S., 80, 305, 309 DeArmond, S. J., 10, 13, 23, 24, 25, 26, 27, 80, 81, 103, 104, 105, 108, 113, 123, 125, 135, 137, 166, 168, 169, 188, 197, 198, 199, 201, 204, 225, 226, 228, 264, 267, 269, 271, 299, 300, 305, 306, 307, 308, 309, 310, 311, 330, 332 Dearmond, S. J., 227, 228 DebBurman, S., 108, 135 DebBurman, S. K., 8, 9, 14, 23, 151, 153, 157, 160, 165, 245, 246, 248, 265, 356, 357, 362 Deboer, T., 81, 228, 311 Dees, C., 199 DeGunzburg, J., 225 Del Bo, B. R., 197 Del Bo, R., 306 Deleage, G., 116, 135 Deleu, C., 325, 330, 332 Delius, H., 198 Delius, J., 23, 166 Della Torre, P., 197 Della Vedova, F., 200 Demaimay, R., 152, 163, 164, 165, 257, 260, 263, 265 Demart, S., 165, 263, 265 Demianova, M., 240, 265 DeMott, D. L., 225 Denayrolles, M., 332 Deng, H., 227 DePace, A. H., 101, 103, 345, 350, 351, 362 Derkach, I. L., 330, 361 Derkatch, I. L., 319, 327, 330, 339, 343, 344, 345, 350, 362 Desbruslais, M., 166, 307

372

AUTHOR INDEX

Deslys, J. P., 165, 167, 198, 200, 263, 265, 267 Deslys, J.-P., 164, 165, 265, 307 Devine-Gage, E., 136 Di Giamberardino, L., 227 Dickenson, A. G., 296, 306 Dickinson, A., 265 Dickinson, A. G., 163, 166, 304, 327, 330 Dickson, D. W., 25, 199, 200, 227, 309 Didichenko, S. A., 364 Diedrich, J. F., 303, 306 Dingwall, W. S., 307 Diomede, L., 197, 200 Diringer, H., 24, 141, 163, 165, 166, 255, 256, 258, 265 Dlouhy, S., 201 Dlouhy, S. R., 25, 26, 105, 169, 198, 200, 201, 227, 228 Doboer, T., 271 Dobson, C. M., 103 Dockter, J., 25, 309 Dodelet, V., 236, 242, 243, 264 Dodelet, V. C., 204, 225 Doel, S. M., 319, 330, 338, 344, 362 Doey, L. J., 22, 166, 305, 307 Doh, u. K., 267 Doh-ura, K., 24, 113, 135, 163, 166 Dohura, K., 104 Dong, A., 22, 135, 165, 197, 305 Doni, R., 197, 306 Donne, D., 136 Donne, D. G., 6, 9, 23, 59, 64, 72, 73, 80, 84, 85, 103, 104, 110, 118, 120, 130, 135, 141, 166, 171, 176, 185, 197, 265, 280, 306 Donnelly, C. A., 21 Dormond, D., 198, 200 Dormont, D., 164, 165, 167, 263, 265, 267, 307 Douglas, M. G., 354, 362 Drake, A. F., 26 Dratz, E. A., 267 Dreosti, I. E., 227 Drillien, R., 317, 330 Drohan, W. N., 271 Drummond, D., 22, 165, 197, 305 Dryden, A. J., 25, 167, 305, 309 Dubinin, N., 362 Duchen, L. W., 286, 306 Dujon, B., 314, 331 Dumpitak, C., 51, 54, 102, 164 Dunker, S., 307

Dunstan, S. P., 21 Dwek, R. A., 39, 45, 53, 81, 137, 168 Dyson, H. J., 23, 80, 82, 103, 105, 135, 166, 197, 201, 265, 306

E Eaglestone, S. S., 336, 362 Eaton, W. A., 362 Ebeling, C., 311 Ebihara, K., 344, 362 Edelbluth, C., 165 Edenhofer, F., 232, 241, 245, 246, 265, 269, 271 Edington, N., 260, 261, 268 Edskes, H. K., 313–329, 331, 361, 362 Ehlers, B., 163, 166, 255, 256, 258, 265 Ehrensperger, F., 80, 136, 198 Eigen, M., 148, 166 Eikelenboom, P., 201 Eklund, C. M., 168 Eliasson, C., 271 Ellis, S. R., 265 Empson, R. M., 307 Endo, T., 30, 37, 39, 53, 81, 125, 135, 137, 168, 206, 225, 299, 306 Engel, J., 263 Engel, W. K., 263 Englander, S. W., 79 Engstrom, U., 331 Equestre, M., 102 Ernst, D., 22, 25, 135, 165, 197, 225, 227, 264, 305, 309 Ernst, M., 80, 198 Ernst, R. R., 79 Erpel, S., 27 Erpel, S. P., 308 Escaig, H. F., 265 Escaig-Haye, F., 225 Essex, J. W., 129, 136 Estibeiro, K., 27, 201, 311 Eurwilaichitr, I., 344, 362

F Fabbrini, G., 102 Fabrizi, C., 200 Falick, A. M., 34, 53 Falls, D. L., 198 Famulok, M., 265, 271 Farlow, M., 25, 199, 309

373

AUTHOR INDEX

Farlow, M. R., 25, 26, 105, 166, 169, 198, 200, 201, 228 Farquhar, C., 166, 255, 256, 258, 265 Farquhar, C. F., 23, 30, 53, 103, 163, 166, 197 Farr-Jones, S., 24, 80, 104, 136, 167 Fasman, G. D., 111, 116, 135 Faucheux, B., 227 Feldmann, C., 23 Fenandez, M.-T., 265 Fenn, J. B., 29, 52 Fenton, W., 267 Ferguson, M. A. J., 34, 45, 53 Ferguson, N. M., 21 Fernandez, M., 240, 266 Ferrari, M., 268 Ferrari, S., 228 Ferrin, T. E., 134, 135 Fersht, A. R., 82, 332 Fesik, S. W., 62, 80 Feuerstein, B., 198, 226 Fields, B. N., 333 Fink, A. L., 129, 135 Fink, G. R., 336, 362 Fisch, E., 271 Fischbach, G. D., 198 Fischer, M., 16, 21, 22, 23, 25, 80, 81, 96, 103, 105, 197, 225, 228, 264, 270, 271, 278, 280, 281, 282, 300, 302, 303, 304, 305, 306, 309, 310, 330 Fischer, R., 80, 136, 198 Fisher, S., 30, 53, 225 Flechsig, E., 270, 306, 310 Fleminger, S., 195, 197 Fletterick, R., 104, 135, 198 Fletterick, R. J., 23, 25, 52, 54, 79, 80, 81, 102, 103, 104, 136, 167, 199, 307, 309 Florio, T., 189, 194, 195, 197 Fluethmann, H., 225 Fogh, J., 167 Foidart, J. M., 271 Forlenza, O. V., 266 Forloni, G., 20, 23, 169, 171–196, 197, 198, 200, 201, 270, 301, 306 Formosa, T. G., 265 Forrest, J., 199, 269 Forrest, J. M., 199 Foster, D., 24, 25, 26, 81, 168, 307, 309, 310, 311, 332 Foster, J., 23, 305 Foster, J. D., 24

Fournier, J. G., 167, 229, 265, 267 Fournier, J.-G., 209, 225, 307 Fowler, N., 24 Frangione, B., 25, 26, 105, 166, 168, 169, 197, 198, 199, 200, 201, 228, 270 Fraser, H., 22, 159, 165, 197, 201, 304, 305, 306, 330 Fraser, P. E., 22, 79, 103, 197, 225, 263, 304, 323, 331, 364 Freeman, I. L., 264 Freeman, S. J., 332 Fridlander, G., 26, 168 Friedhuber, 198 Friedlander, G., 227 Fritch, W., 268 Frolova, L., 333, 364 Frolova, N. S., 362 Fujimoto, K., 265 Fujioka, H., 105, 227 Fujita, K., 362 Futerman, A. H., 227

G Gabizon, R., 9, 19, 20, 23, 24, 26, 27, 81, 100, 103, 104, 105, 137, 166, 168, 169, 198, 231, 255, 264, 265, 271, 310 Gabriel, J. M., 80, 135, 177, 198 Gabriel, J.-M., 136, 307 Gadjusek, D. C., 8, 23, 146, 166, 167 Gahali, I., 137 Gajdusek, D. C., 16, 23, 26, 55, 79, 81, 104, 137, 168, 197, 199, 200, 227, 304, 308 Galceran, J., 269 Galkin, A., 361 Gall, S., 226 Gallo, G., 198 Gambetti, P., 22, 25, 27, 103, 104, 105, 136, 137, 167, 168, 169, 199, 221, 225, 227, 228, 264, 308, 309, 310 Gambliel, H. A., 225 Gane, P. J., 250, 271 Garcia, A. J., 268 Garcia, F. L., 6, 23, 27 Garcia-Hernandez, M., 239, 240, 265 Garini, P., 268 Garnier, J., 111, 116, 135, 136 Gasset, M., 8, 23, 25, 81, 84, 102, 103, 104, 109, 113, 115, 120, 135, 136, 167, 175, 180, 182, 185, 198, 199, 309 Gauczynski, S., 229–262

374

AUTHOR INDEX

Gaus, S. E., 81, 228, 271, 311 Gebert, R., 80, 104, 362 Geelen, J. L., 200 Gekko, K., 248, 265 Gelderblom, H., 165 Gelinas, R.E., 304 Genbauffe, F., 332 Georgopoulos, C., 361 Geourjon, C., 116, 135 Gerbert, R., 331 German, T. L., 199 Geroni, C., 263 Gething, M. J., 245, 265 Geuze, H. J., 102 Ghetti, B., 25, 26, 105, 166, 167, 169, 172, 174, 175, 184, 197, 198, 199, 200, 201, 225, 227, 228, 305, 308, 309 Ghibaudi, E., 197, 200, 201, 306 Ghirelli, C., 264 Ghiso, G., 198 Ghiso, J., 26, 198, 199, 201, 228 Giaccone, G., 26, 105, 146, 166, 169, 174, 197, 198, 199, 200, 201, 227, 228, 270, 306 Gianni, A. M., 265 Gianni, L., 261, 265 Gibbs, C. J., 81, 193, 198 Gibbs, C. J. J., 16, 23, 304 Gibbs, C. J. Jr., 26, 104, 137, 168, 197, 200, 227 Gibrat, J. F., 116, 135 Gibson, B. W., 33, 34, 36, 39, 41, 43, 53, 105, 137 Gibson, P. H., 169 Gielkens, A. L. J., 21 Giese, A., 10, 22, 23, 25, 79, 103, 167, 197, 225, 226, 263, 304 Gilbert, I. H., 168 Gilch, S., 27, 82, 105 Gill, A. C., 263 Gilman, A. G., 225 Gioia, B., 263 Giuliani, F. C., 263 Glabe, C. G., 200 Glassmith, L. L., 22 Glenn, T., 310 Glenner, G. G., 168, 200, 253, 265 Glockshuber, R., 26, 61, 78, 79, 80, 81, 83–102, 103, 104, 105, 109, 126, 134, 135, 136, 137, 140, 148, 153, 154, 166, 168, 198, 199, 200, 213, 226, 262, 270, 283, 307, 309

Glover, J. R., 8, 23, 101, 103, 269, 328, 331, 346, 347, 348, 349, 352, 355, 358, 362, 363 Goedert, M., 198 Goethals, M., 200 Goetz, J., 270 Golbe, L. I., 227 Golbik, R., 82 Golden, G. T., 199, 227 Goldfarb, L. G., 18, 23, 100, 103, 167, 176, 198, 283, 304, 306, 308 Goldin, L., 306 Goldmann, W., 15, 16, 23, 24, 166 Goller, N. L., 224 Goloubinoff, P., 363 Gomi, H., 252, 265 Gonatas, N., 25, 199, 226 Gonzales, M., 197 Gonzalez, D. P., 268 Goodin, D. B., 82, 105, 201, 228 Goodman, P. A., 27, 305, 311 Goodsir, C. M., 24, 166, 226 Gordon, H., 165 Gorevic, P., 201 Gorla, S., 200 Gorodinsky, A., 211, 221, 225, 266 Gottesman, S., 364 Gotz, J., 310 Gowland, I., 22, 80, 166, 228, 305, 306, 307, 311 Graham, C. H., 226 Graham, K., 227, 309 Graner, E., 254, 255, 266, 268 Graves, F. M., 362 Gray, F., 167, 308 Gray, P. C., 168 Gray, V. T., 331, 362 Gready, J. E., 128, 137 Greenberg, B. D., 268 Greig, A., 307 Greiner, R., 80, 103 Greiner, R. A., 264, 305 Greiner, R.-A., 22, 330 Griffin, J., 264 Griffith, J., 338, 339, 348, 362 Griffith, J. S., 3, 6, 23, 55, 56, 74, 80, 83, 103, 139, 146, 166, 327, 331 Griffith, O. H., 21, 24, 166, 263 Grimaldi, M., 197 Grimes, G. W., 327, 331 Gritzmacher, C. A., 267

AUTHOR INDEX

Groschup, M., 271 Groschup, M. H., 23, 24, 271, 305 Gross, H., 80, 104, 331, 362 Groth, D., 23, 24, 25, 26, 30, 37, 39, 53, 80, 81, 102, 103, 104, 105, 135, 136, 137, 166, 167, 168, 169, 197, 198, 199, 225, 226, 227, 265, 306, 307, 309, 310, 311, 332 Groth, D. F., 24, 30, 31, 53, 79, 135, 168, 200, 304, 307 Grzesiek, S., 61, 79 Guarnieri, M., 224 Gunning, R. F., 27 Güntert, P., 63, 79, 80 Gyuris, T., 270

H Haase, A. T., 306 Hadlow, W. J., 168 Hafiz, F., 22, 264 Haga, T., 27 Haggqvist, B., 328, 331 Haghighar, A., 264 Haig, D. A., 21, 79, 102, 330 Haïk, S., 187, 198, 200 Hainfellner, J., 199, 227 Haist, I., 23, 24 Halimi, M., 23, 103, 264, 265 Hall, S., 226 Halladay, J., 361 Halliday, W. G., 24, 226 Haltia, M., 198 Han, D., 27 Hancock, R. D., 27 Hangartner, C., 270, 310 Hanover, J. A., 331 Hanson, R. P., 200 Haraguchi, T., 30, 53, 206, 225 Harding, A. E., 305, 306, 308 Harper, J., 165 Harris, D., 305 Harris, D. A., 10, 19, 22, 24, 49, 53, 100, 103, 104, 168, 169, 177, 179, 186, 198, 199, 200, 203–224, 225, 226, 227, 228, 231, 241, 266, 270 Harrison, P. M., 8, 23, 108, 135, 226, 268, 308 Hartl, U., 271 Harvey, D. J., 39, 53, 81, 137, 168 Hasegawa, S., 227, 270 Hässig, R., 227

375

Haswell, S., 22 Haswell, S. J., 22, 264 Hauw, J. J., 167, 199, 227, 267, 308 Hauw, J.-J., 307 Hawkins, P. N., 103, 168 Hawthorne, D. C., 341, 362 Hay, B., 19, 23, 218, 226, 284, 306 Hayes, S. F., 22, 135, 165, 197, 305 Hecker, R., 41, 42, 43, 45, 47, 53, 227 Hedge, R. S., 95, 103, 186, 198, 284, 285, 286, 306 Heffner-Lauc, M., 199 Hegde, R. S., 10, 11, 19, 20, 23, 58, 80, 146, 166, 218, 221, 226 Hegyi, I., 25, 270, 309, 310 Heimark, R. L., 270 Heinrich, C., 169, 201, 226, 268, 308, 310 Heller, J., 79, 80, 180, 198 Helms, C., 332 Henner, D., 104 Hennion, R. M., 168 Hermann, L. M., 130, 135 Hermanowski-Vosatka, A., 226 Herms, J., 197, 209, 226 Herms, J. W., 22, 79, 103, 197, 225, 263, 275, 304, 306, 307 Herrmann, L. M., 9, 23, 93, 103, 150, 154, 166 Hess, H. H., 196 Heuser, J. E., 227, 270 Hietala, S. O., 168 Hill, A. F., 22, 24, 80, 103, 105, 136, 143, 151, 156, 162, 165, 166, 169, 201, 228, 291, 293, 294, 295, 298, 305, 306, 307, 311 Hillenkamp, F., 29, 52 Hillerton, J. E., 21 Hillner, P., 103, 362 Hilmert, H., 165 Hinek, A., 238, 266 Hnatowich, M., 269 Hockenbery, D., 254, 266 Hoffmann, A., 266 Hofman, A., 27, 201, 311 Hofrichter, J., 349, 362 Hogan, V., 268 Hoinville, L. J., 21 Holscher, C., 16, 23, 145, 166 Hölscher, C., 180, 198 Holtzman, D. M., 26, 135, 198, 227 Holzemann, G., 201

376

AUTHOR INDEX

Homans, S. W., 45, 53 Hong, B. L., 263 Hood, L., 30, 33, 36, 39, 53, 105, 137, 225 Hood, L. E., 27, 30, 52, 53, 81, 82, 105, 137, 167, 169, 200, 226, 228, 268, 269, 308 Hoope, P. C., 25 Hooper, M. L., 267, 307 Hooper, N. M., 225 Hope, J., 4, 20, 22, 23, 24, 25, 26, 30, 36, 39, 53, 84, 102, 103, 104, 135, 141, 146, 148, 165, 166, 168, 169, 190, 192, 197, 198, 224, 226, 228, 267, 305, 307, 308, 310 Hoppe, P. C., 168, 309, 332 Hori-i, A., 199, 226 Hori-i, H., 25 Horiuchi, M., 8, 9, 14, 17, 24, 83, 103, 108, 123, 135, 136, 140, 149, 151, 153, 154, 155, 156, 157, 160, 165, 166, 264 Hornemann, S., 26, 61, 79, 80, 81, 84, 85, 86, 87, 91, 92, 102, 103, 104, 109, 134, 135, 136, 137, 141, 148, 166, 168, 198, 199, 200, 262, 270, 309 Hornshaw, M. P., 10, 24, 50, 53, 177, 178, 198, 205, 226, 281, 307 Horwich, A. L., 364 Hosie, B. D., 307 Hosszu, L., 103, 136 Hosszu, L. L., 24, 72, 80, 86, 87, 91, 103, 166 Hosszu, L. L. P., 131, 136 Houghten, R. A., 82, 136 Houtani, T., 227, 270, 308, 310 Howley, P. M., 333 Hsiao, K., 26, 30, 53, 81, 228, 274, 283, 286, 287, 301, 307, 308 Hsiao, K. K., 20, 24, 81, 113, 136, 225, 302, 307 Hsu, D. K., 272 Huang, C. C., 135 Huang, Z., 25, 52, 54, 67, 80, 81, 96, 102, 103, 104, 111, 112, 115, 116, 135, 136, 167, 199, 281, 307, 309 Huber, M. T., 49, 53, 225 Huber, R., 79 Hughes, J. T., 25, 167, 309 Hughes, K., 23 Hughes, R. C., 237, 263 Hughes, S., 23 Hundt, C., 229–262 Hunter, D. D., 266

Hunter, G. D., 199 Hunter, I., 263 Hunter, N., 15, 16, 23, 24, 166, 293, 307 Hunziker, P., 50, 53 Hutchinson, W. L., 103 Hyun, W., 228 Hyun, W. C., 198, 226

I Ichimiya, Y., 198 Iizuka, R., 24, 198 Ikeda, T., 265 Ikonen, E., 212, 227 Imahori, K., 364 Imperiali, B., 13, 25, 299, 308 Inaba, H., 268 Inge-Vechtomov, S., 333, 341, 362, 364 Inge-Vechtomov, S. G., 264, 330, 361, 362, 364 Ingedoh, A., 29, 52 Ingrosso, L., 163, 166, 167, 257, 258, 266, 267, 271 Inouye, H., 167, 181, 198, 199 Iqbal, K., 167, 199 Ironside, J., 22, 25, 103, 104, 165, 168, 169, 200, 270, 306, 308, 310 Ironside, J. W., 22, 27, 165, 197, 201, 305, 311 Isenmann, S., 21, 197, 304 Itoh, T., 265 Itohara, S., 265 Itri, V., 81, 168 Iu, O., 362 Ivanova, 213 Iwahashi, H., 362 Iwaki, T., 166 Iwamatsu, A., 270 Izquierdo, I., 266

J Jackers, P., 240, 264, 266 Jackson, G. S., 9, 24, 80, 86, 87, 91, 92, 93, 94, 95, 96, 101, 103, 105, 130, 136, 148, 154, 166, 169, 201 Jaegly, A., 167, 267, 307 Jaffe, H., 104 Jaffe, R., 264 James, J. L., 135

AUTHOR INDEX

James, J. T., 136 James, T. L., 23, 24, 59, 71, 80, 82, 84, 85, 103, 104, 105, 110, 116, 118, 119, 120, 126, 128, 130, 136, 137, 166, 167, 169, 197, 201, 265, 266, 306 Jamieson, E., 25, 308 Jankowski, R., 103 Jansen, V. A., 167 Jansen, V. A. A., 308 Jardetsky, O., 136 Jares, P., 264 Jarrett, J. T., 8, 24, 84, 101, 104, 146, 147, 148, 166, 348, 349, 362 Jarvis, L. E., 135 Jaye, M., 271 Jefferys, J. G., 225, 228 Jefferys, J. G. R., 80, 225, 264, 275, 300, 305, 307, 311 Jeffrey, M., 6, 24, 27, 166, 216, 226 Jen, A., 226 Jensen, M., 167 Jin, T., 228 Jing, Y., 228 Jobling, M. F., 193, 198 Jobling, M. R., 135 Johnson, C. M., 82 Johnson, C. T., 27 Johnson, F. A., 198 Johnson, K. H., 332 Johnson, R. T., 193, 198 Johnston, A., 226, 308 Joiner, S., 105, 166, 169, 201, 307 Jones, C. K., 227 Jones, E. W., 330 Jones, I. M., 22, 264, 271 Jones, K. M., 288, 309, 332, 363 Joris, B., 264 Joseph, C., 103, 136 Ju, W. K., 198 Jucker, M., 266 Juliano, M. A., 266 Julien, J., 22, 103 Jushage, K., 309

K Kaczkowski, J., 268 Kagen, B. L., 199 Kahana, I., 23, 103 Kaneda, Y., 266, 270

377

Kaneko, K., 10, 19, 20, 24, 27, 77, 80, 82, 98, 101, 104, 136, 137, 155, 156, 162, 166, 167, 184, 198, 221, 226, 247, 261, 266, 269 Kanelo, K., 201 Kanoush, R., 228 Kaplan, B., 173, 198 Karas, M., 29, 52 Karger, B., 331 Karunaratne, A., 226, 268, 308 Kascsak, R., 168, 169, 200, 270 Kascsak, R. J., 11, 24, 146, 160, 161, 166, 167, 168, 169, 200, 224, 270 Katamine, S., 226, 227, 268, 270, 308, 309, 310 Kataoka, Y., 227, 270, 310 Katoh, A., 265 Katzenstein, G. E., 264 Katzenstien, G. E., 22 Kaufman, M. H., 226, 307 Kaufmann, R., 51, 54, 104, 167 Kawai, R., 355, 362 Kawai, S., 269 Kazmirski, S., 115, 134, 136 Kazmirski, S. L., 113, 114, 136 Keller, B. U., 306 Kellings, K., 168, 262, 266 Kelly, J. W., 92, 102, 104 Kelly, S. M., 23, 198 Kelve, M., 199, 269 Kenaga, L., 25, 167, 199, 226 Kenney, J., 24, 103, 136, 166 Kent, S. B., 29, 52, 200 Kent, S. B. H., 30, 52, 81, 167 Keppel, E., 239, 240, 266 Kessel, M., 355, 362 Khakoo, A. Y., 240, 264 Khana, M., 225 Kibbey, M. C., 254, 266 Kiersch, D., 51, 54 Kikuchi, A., 362 Kikuchi, Y., 342, 344, 362 Kikuno, R., 269 Kim, B., 362 Kim, J. I., 196, 198 Kim, Y. S., 198, 306 Kimberlin, R., 30, 53 Kimberlin, R. H., 2, 9, 23, 24, 25, 26, 103, 105, 137, 162, 163, 165, 166, 167, 168, 199, 255, 256, 266

378

AUTHOR INDEX

King, C. Y., 79, 80, 101, 104, 346, 347, 362 King, C.-T., 328, 331 King, J., 137 Kingsbury, D. T., 225 Kinoshita, K., 239, 240, 266, 270 Kinsbury, D. T., 305 Kirsch, D., 104, 167, 269 Kirschner, D. A., 167, 181, 198, 199 Kish, S. J., 200 Kisilevsky, R., 26, 268, 323, 331 Kisselev, L., 333, 364 Kitamoto, T., 17, 18, 20, 24, 25, 100, 104, 135, 167, 184, 198, 229, 267 Kitamura, M., 268 Klein, M. A., 21, 25, 304, 306, 309 Klein, T. R., 51, 54, 101, 104, 162, 167 Kleinman, H. K., 266 Kline, A. D., 79 Klunk, W. E., 347, 362 Kneller, D. G., 111, 116, 136 Knipe, D. M., 333 Kobata, A., 30, 37, 39, 53, 81, 135, 137, 168, 225, 306 Kobayashi, G. S., 268 Kobayashi, Y., 21, 197 Kobayshi, Y., 304 Kochneva-Pervakhova, N. V., 350, 351, 362 Kocisko, D. A., 3, 8, 9, 10, 12, 15, 20, 21, 22, 24, 25, 79, 102, 104, 108, 109, 125, 135, 136, 140, 143, 144, 150, 151, 153, 156, 159, 164, 165, 167, 168, 328, 330, 331 Kocsis, E., 362 Koehler, R., 26, 309, 310 Kohler, R., 26, 168 Kolbert, A. C., 80, 198 Koliatsos, V. E., 224 Komatsu, Y., 362 Kon, A. A., 197 Kondig, J. P., 267 Koo, E. H., 254, 267 Kopp, N., 25, 167 Koradi, R., 63, 80, 104 Korsmeyer, S. J., 253, 266, 269, 271 Korth, C., 78, 80, 103, 121, 123, 129, 136, 166, 184, 198 Kosic-Smithers, J., 364 Kowal, A. S., 23, 103, 331, 362, 363 Krajewski, S., 254, 267 Kraulis, P. J., 134, 136 Kreeger, T. J., 167

Kretschmar, H., 80 Kretzschmar, H., 22, 25, 79, 103, 136, 167, 198, 225, 226, 263, 304 Kretzschmar, H. A., 22, 23, 197, 204, 225, 226, 271, 274, 302, 304, 306, 307 Kristensson, K., 185, 198, 212, 226 Krobitsch, S., 356, 357, 362 Kruck, T., 22, 79, 103, 197, 225, 263, 304 Krutzsch, H. C., 264 Kshet, I., 193, 198 Kuczius, T., 13, 24 Kudo, A., 269 Kühl, U., 267 Kuhn, R. J., 271 Kumar, A., 198 Kumar, J., 361 Kunau, W.-H., 327, 331 Kuno, J., 309 Kunz, B., 50, 53, 193, 198, 261, 267, 301, 307 Kurokawa, K., 309 Kurschner, C., 236, 253, 267 Kurzhalla, T. V., 199 Kushnirov, V. V., 310, 330, 332, 342, 343, 344, 351, 357, 358, 362, 363, 364 Kyrpides, N., 269

L Lacroute, F., 314, 316, 317, 319, 329, 330, 331, 332 Ladogana, A., 162, 166, 167, 255, 256, 258, 266, 267, 271 Laing, J. G., 237, 267 Lakey, J. H., 24, 50, 53, 198, 226 Laloraya, S., 361 Lamoury, F., 263 Landon, M., 102, 224 Landowski, T. H., 240, 267 Landsbury, P. T., 330 Lang, Y., 80, 225, 264, 305, 330 Langen, H., 363 Langeveld, J. P., 198 Langridge, R., 135, 136 Lansbury, P. T., 79, 102, 135, 136, 165, 167, 315, 331 Lansbury, P. T. Jr., 8, 21, 22, 24, 25, 84, 101, 104, 136, 146, 147, 148, 164, 165, 166, 167, 168, 347, 349, 352, 361, 362 Lansen, J., 169

AUTHOR INDEX

Lantos, P., 80, 166, 307 Lantos, P. L., 22, 305, 306 Larsen, C. T., 167 Larsen, R., 80, 198 Lasen, J., 270 Laskowski, R. A., 136 Lasmezas, C., 164, 165, 265 Lasmézas, C., 198 Lasmezas, C. I., 162, 165, 167, 263, 270, 301, 307 Lasmézas, C. I., 200, 229, 262, 265, 267, 269 Last, A. M., 50, 53 Laszlo, L., 26, 102, 201, 224, 227, 228, 270 Lauro, G. M., 200 Lawrence, J. D., 80 Lazarini, F., 165, 265 Lazarow, P. B., 327, 331 Lazzarini, A., 227 Le Goff, X., 364 Le Guellec, R., 364 Lea, H. Z., 332 Leach, M., 308 Leader, W. M., 269 Leal, S., 308 LeBlanc, A., 308 Leclerc, A., 263 Lee, C., 272 Lee, I. Y., 226, 268, 308 Lee, R. J., 104 Lefkowitz, R. J., 269 LeGoff, X., 333 LeGuellec, R., 333 Lehmann, S., 10, 19, 22, 24, 100, 103, 104, 186, 199, 207, 208, 209, 213, 216, 219, 225, 226, 227, 266 Lele, P., 226 Lennox, A., 227 Lenzi, G. L., 102 Lesko, A., 227, 270 Lesot, H., 237, 239, 267 Leucht, C., 229–262 Leupold, U., 341, 362 Levin, J. M., 116, 136 Levy, E., 168 Lewitin, E., 361 Li, L., 361, 362 Liang, Y., 226, 268 Liautard, J. P., 244, 267 Lieberburg, I., 23 Lieberburg, S. B., 306

379

Liebman, S. W., 264, 330, 361, 362 Liemann, S., 78, 80, 86, 87, 90, 96, 97, 99, 104, 105, 126, 136, 226, 283, 307 Lievens, P. M. J., 175, 199, 200 Lievens, P. M.-J., 169, 270 Lin, M. C., 187, 199 Lindquist, S., 9, 23, 24, 95, 96, 103, 104, 135, 165, 208, 222, 226, 265, 269, 331, 335–361, 362, 363 Lindquist, S. L., 264, 361, 363 Lingappa, V. R., 23, 24, 80, 103, 166, 198, 225, 226, 228, 284, 306, 311 Lins, L., 198, 200 Liotta, L., 271 Liotta, L. A., 269 Lipp, H., 80 Lipp, H. P., 264, 305, 330 Lipp, H.-P., 225 Lisanti, M. P., 211, 226 Lithgow, T., 254, 267 Liu, A., 27, 64, 73, 80, 82, 105, 137, 184, 199, 201 Liu, F. T., 267, 272 Liu, H., 5, 6, 7, 24, 59, 71, 72, 74, 75, 77, 80, 104, 118, 128, 136, 141, 155, 167 Liu, J., 23 Liu, J. J., 269, 338, 341, 345, 350, 351, 357, 362, 363 Liu, J.-J., 103, 331 Liu, Y., 130, 136 Livshits, T., 201 Livshits, T. L., 82, 137 Lledo, P., 275, 307 Lledo, P. M., 229, 232, 267 Llinas, M., 24, 80, 136, 167 Lloyd, D. H., 135, 198 Lofthouse, R., 305, 308 Loftus, B., 77, 80 Loo, J. A., 50, 53 Lopez Garcia, F., 105, 137, 171, 199, 201 Lopez, C. D., 19, 24, 218, 226, 228, 311 Lopez-Buesa, P., 363 López-García, F., 73, 78, 80, 82 Lopez-Hoyo, N., 363 Lopez-Ribot, J. L., 239, 240, 241, 267 Lovett, M., 305 Low, M. G., 208, 226 Low, P. S., 104 Lowenstein, D. H., 288, 307 Lowery, D. E., 268

380

AUTHOR INDEX

Lu, Z. Y., 196 Lucca, E., 197, 200 Ludwig, G. V., 239, 240, 241, 267 Lugaresi, A., 167 Lugaresi, E., 22, 27, 103, 137, 169, 308, 310 Luginbühl, P., 63, 80 Luhrs, T., 27, 137, 201 Lührs, T., 82, 105 Lund, P. M., 331, 336, 363 Lundberg, K. M., 7, 24 Lycke, E., 30, 53, 225 Lysek, D. A., 80

M Ma, J., 9, 24, 95, 96, 104, 208, 222, 226 MacArthur, M. W., 136 Macchi, G., 102, 268 Mackenzie, I. R., 200 MacLeod, N., 226, 308 Maddelein, M.-L., 313–329, 331 Madlung, A., 22, 79, 197, 225, 226, 263, 304 Madore, N., 212, 226 Madung, A., 103 Maeda, N., 271 Magasanik, B., 316, 317, 318, 319, 330, 331 Magnifico, A., 263, 264 Maher, F., 198 Malesani, P., 197, 200 Malinoff, H. L., 237, 239, 267 Man-A-Hing, W. K., 201 Manetto, V., 308, 309 Mann, M., 29, 52 Manson, J., 22, 79, 103, 197, 200, 204, 225, 226, 263, 287, 288, 304, 307 Manson, J. C., 17, 20, 25, 81, 204, 205, 228, 229, 232, 267, 271, 275, 278, 307, 308, 311 Manuelidis, E. E., 164, 199 Manuelidis, L., 164, 196, 199, 260, 261, 268 Maras, B., 102, 105 Marino, S., 21, 197 Marinone, M. G., 268 Markley, J. L., 82 Marmostein, A. D., 21 Marquesee, S., 80 Marqusee, S., 24, 136, 167 Marrari, M. A., 200 Marsahall, S. T., 305 Marsh, R., 167, 268

Marsh, R. F., 2, 11, 12, 21, 25, 79, 142, 145, 146, 160, 161, 164, 165, 173, 185, 197, 199, 200, 215, 224 Marthin, T., 332 Martin, G. R., 271 Martin, R., 197 Martin, T., 23 Martinez, J. P., 267 Martins, V. R., 235, 236, 266, 268 Masel, J., 145, 148, 167, 274, 308 Masison, D. C., 319, 321, 322, 324, 327, 331 Mason, J., 308 Mason, J. C., 308 Masters, C. L., 135, 198, 253, 268 Mastrangelo, P., 226, 268, 308 Mastrianni, F., 103 Mastrianni, J., 26, 27, 81, 105, 137, 169, 271, 310 Mastrianni, J. A., 23, 80, 198, 226, 306 Masullo, C., 167, 260, 268, 269, 271 Matsunaga, Y., 136 Matthiesen, U., 51, 54, 102, 164 Maurizi, M. R., 362, 364 MaWhinney, S., 21 Mayer, R. J., 102, 224 Mayne, L., 79 McArthur, R. A., 169, 270 McBride, P., 226, 271, 307 McBride, P. A., 24, 25, 81, 166, 197, 226, 228, 308 McCardle, L., 22, 165, 197, 305 McCarty, C. W., 167 McCloskey, J. A., 29, 52 McConnell, I., 22, 25, 165, 197, 226, 267, 304, 305, 307, 308 McCready, S. J., 330, 338, 362, 363 McDermott, J. R., 24, 50, 53, 198, 226, 307 McDonald, B. L., 268 McFadden, P. N., 50, 53 McHattie, S., 260, 261, 268 McHolland, L. E., 168 McKenzie, D., 143, 164, 167, 260, 268 McKenzie, D. I., 79, 164 McKindly, M. P., 311 McKinley, M. P., 5, 6, 25, 26, 27, 30, 31, 52, 53, 79, 81, 135, 146, 167, 168, 185, 199, 200, 216, 220, 226, 280, 304, 307 McLachlan, D. C., 227 McLaughlin, C. S., 330, 361, 363 McLean, C., 198

AUTHOR INDEX

McLean, M. J., 309 McNutt, P., 243, 270 Meads, J., 22, 80, 103, 165, 228, 305, 306, 311 Mecham, R. P., 239, 240, 266, 268 Medoff, G., 259, 268 Medori, R., 283, 306, 308 Meesters, E., 199 Mehlhorn, I., 23, 25, 80, 81, 84, 87, 93, 102, 103, 104, 105, 135, 136, 137, 166, 167, 169, 197, 199, 265, 269, 306, 309 Meikle, V., 306 Meikle, V. M. H., 330 Meiner, Z., 23, 26, 103, 137, 168, 265 Meinhardt, A., 199 Melnick, M. B., 239, 240, 268 Melton, D., 308 Melton, D. W., 25, 226, 268, 308 Menard, S., 263, 264 Mendez, E., 169, 200, 270 Meng, C. K., 29, 52 Mercadante, A. F., 266, 268 Merenda, C. A., 311 Mering, C. v., 310 Merlie, J. P., 266 Merlini, G., 261, 265, 268 Merz, P. A., 24, 141, 166, 167, 182, 199 Messerle, B. A., 81 Meyer, D. I., 354, 361 Meyer, N., 266 Meyer, R. K., 167 Migheli, A., 189, 199 Mikol, J., 22, 103, 167, 308 Miller, M. W., 157, 167, 168 Millhauser, G. L., 24 Milliman, C., 266 Milliman, C. L., 269 Millson, G., 185, 199 Millson, G. C., 26, 105, 137, 168 Milne, J. S., 79 Minamiya, Y., 268 Minetti, A., 266 Minoletti, F., 266 Miranker, A. D., 364 Miravalle, L., 173, 199 Mirels, L. F., 225 Mirenda, C., 25, 26, 81, 168, 266, 309, 310, 332 Mirenda, C. A., 27 Mirwald, J., 167

381

Mirzabekov, T., 199 Mitchell, A. P., 317, 318, 331 Mitrova, E., 306 Miura, T., 10, 25, 178, 179, 199, 205, 226 Miyamoto, T., 227, 270, 309 Miyazono, M., 267 Mobley, W. C., 26, 197, 225, 227 Mochi, M., 308 Moffat, B., 104 Moffat, B. A., 61, 81 Mogk, A., 352, 355, 363 Mohn, C. M., 195, 197 Molinari, A., 200 Monari, L., 146, 161, 167, 300, 308 Montagna, P., 27, 137, 167, 169, 308, 309, 310 Monteagudo, C., 267 Monticelli, L., 200 Monzani, E., 23, 197 Moore, L., 307 Moore, R., 276, 308 Moore, R. C., 15, 25, 204, 208, 226, 230, 232, 233, 234, 268, 276, 277, 278, 283, 296, 308 Morgan, J. I., 236, 253, 267 Morillas, M., 102, 104, 105, 124, 136, 150, 167 Morimoto, R., 361 Moriuchi, R., 227, 270, 309, 310 Moriyama, H., 313–329 Morris, R., 226 Morton, L. J., 103 Morton, L. J. D., 23, 30, 53, 166 Moser, M., 23, 80, 81, 103, 136, 198, 204, 226, 228, 271, 303, 306, 308 Moslehi, J. J., 363 Moss, D. S., 136 Mössner, E., 105 Mototani, H., 199, 226 Motoyama, S., 254, 268 Moulder, K., 225, 266 Moulder, K. L., 227, 270 Moya, K. L., 227 Mucchiano, G., 331, 332 Mucke, L., 25, 200, 309 Mudry, M., 271 Muileman, I. H., 21 Müller, K. M., 305 Muller, W. E., 199, 269 Müller, W. E., 195, 199

382

AUTHOR INDEX

Mulqueen, O. U., 26, 169, 310 Multhaup, G., 23, 30, 53, 103, 166, 268 Mumby, S. M., 225 Mumenthaler, C., 80 Mundy, C. J., 361 Mundy, C. R., 332, 364 Muramoto, T., 4, 7, 25, 146, 167, 169, 175, 180, 199, 201, 267, 281, 282, 287, 308, 310 Murata, M., 185, 199 Myers, J. K., 90, 104 Myers, R. M., 24, 226, 228, 311

N Nadal, G. B., 269 Nakahara, D. H., 225 Nakamura, Y., 344, 362 Nakaoke, R., 227, 308, 309 Nakatani, A., 227, 270, 309 Nakatani, T., 310 Nandan, S., 21, 79, 102, 135, 164, 330 Nandi, P. K., 262, 268 Narindrasorasak, S., 254, 268 Narwa, R., 186, 199, 216, 218, 227 Naslavsky, N., 211, 221, 227 Naslund, J., 331 Neary, K., 22, 225 Nelson, J., 361 Nelson, R. J., 360, 363 Neri, D., 61, 81 Neto, V. M., 268 Neumann, M., 271 Neve, R. L., 266 Newman, G. P., 356, 358, 359, 363 Newnam, G., 361 Newnam, G. P., 361 Nguyen, H. O., 169, 201 Nguyen, H. O. B., 310 Nguyen, H.-O. B., 310 Nguyen, J., 25, 81, 102, 104, 109, 113, 120, 135, 136, 167, 201, 309 Nguyen, J. T., 82, 137, 141, 167, 180, 183, 184, 199 Nguyen, O., 26, 310 Nicolet, C., 363 Nicoll, R. A., 267, 307 Nierras, C. R., 330, 332, 362, 363 Nilsson, P., 167 Ninchak-Casey, A., 23, 135, 306 Nishida, N., 205, 227, 270, 277, 308, 309, 310

Nishio, J., 22, 135, 165 Nixon, R., 201, 228, 271 Nochlin, D., 200 Noda, T., 227, 270, 309 Noll, E., 268 Nordstedt, C., 331 Nowak, M. A., 167, 308 Nunez, G., 266

O O’Conner, S. E., 13, 25 O’Connor, S. E., 299, 308 O’Dowd, B. F., 253, 269 O’Rourke, K. I., 168 Obata, S., 270 Ochieng, J., 237, 268 Oesch, B., 23, 30, 52, 53, 56, 57, 79, 80, 81, 103, 135, 136, 141, 143, 166, 167, 198, 226, 228, 236, 250, 251, 252, 268, 269, 271, 304, 306, 308 Ogawa, J., 268 Ohba, M., 360, 363 Okada, H., 227, 270 Okazaki, M., 243, 269 Olander, D., 167 Oldstone, M. B. A., 25, 200, 309 Olofsson, S., 30, 53, 225 Oltvai, Z. N., 253, 269 Ono, B., 264, 361 Orso, O., 197 Osguthorpe, D. J., 135 Osherovich, L. Z., 104, 363 Ott, J., 307, 308 Otting, G., 81 Ou, S., 271 Ouzonis, C., 239, 240, 269 Ovadia, H., 198 Owen, F., 283, 286, 305, 307, 308 Ozel, M., 165

P Pace, C. N., 104 Palladini, 185 Palmer, M. S., 15, 22, 25, 26, 80, 159, 167, 225, 228, 264, 305, 306, 309, 311 Palmiter, R. D., 304 Pan, K., 24, 135, 166 Pan, K. M., 25, 59, 65, 81, 84, 102, 103, 104, 108, 109, 136, 168, 171, 178, 199, 309

AUTHOR INDEX

Pan, K.-M., 6, 25, 31, 41, 42, 43, 45, 47, 52, 53, 54, 141, 167, 227, 274, 309, 332 Papini, M. C., 227 Paramithioris, E., 264 Parchi, P., 11, 12, 13, 22, 25, 27, 103, 104, 137, 161, 167, 169, 173, 175, 199, 222, 225, 227, 264, 297, 300, 308, 309, 310 Parchment, O. G., 129, 136 Pardo, C. A., 224 Park, L., 267 Parsell, D. A., 353, 354, 355, 363 Passerini, F., 197, 200, 201 Pasternak, S. H., 226, 268, 308 Patino, M. M., 23, 103, 247, 269, 331, 339, 341, 343, 344, 347, 356, 357, 358, 361, 362, 363 Pattison, I. H., 288, 309 Patton, J. G., 252, 269 Pauly, P. C., 179, 199, 206, 209, 211, 227 Paushkin, S. V., 328, 332, 339, 341, 343, 344, 348, 356, 357, 363 Pavlov, Y. I., 361 Payne, W. N., 197 Pearson, G., 305 Pedersen, P. L., 137 Pedley, T. A., 185, 201 Pedrotti, B., 200 Pekny, M., 271 Penco, S., 263 Peng, J., 63, 65, 81 Pepys, M. B., 103, 168, 364 Peranen, J., 199 Peressini, E., 197 Perett, S., 82 Peretz, D., 21, 24, 82, 121, 123, 129, 136, 166, 169, 201, 263, 310 Perez, M., 192, 199 Perfetti, V., 268 Pergami, P., 84, 104 Pergande, G., 199, 269 Pergrande, G., 199 Peri, E., 169, 270 Perini, F., 198 Perlman, S. L., 227 Permanne, B., 168 Perovic, S., 192, 199, 260, 261, 269 Perrett, S., 321, 332 Perrier, V., 163, 167, 257, 261, 269 Perrimon, N., 268 Perry, B. J., 272 Perry, G., 225

383

Perry, L., 22 Perry, L. L., 225 Perutz, M., 332 Pervushin, K., 79, 81 Pesole, G., 263 Petersen, R., 105, 137, 169 Petersen, R. B., 19, 25, 100, 104, 137, 167, 168, 199, 213, 225, 227, 228, 308 Peterson, R. B., 105, 309 Peterson-Torchia, M., 305 Petraroli, R., 102 Petromilli, 308 Pettegrew, J. W., 362 Peyrin, J. M., 167, 195, 198, 200, 267 Peyrin, J.-M., 307 Pfaff, E., 305 Pfund, C., 360, 363 Phan, U. T., 102 Philippe, M., 333, 364 Piccardo, P., 17, 25, 173, 175, 198, 199, 200, 201, 222, 225, 227, 305 Pike, C. J., 193, 200 Pilkuhn, S., 24, 201, 226, 228, 266, 271 Pillot, T., 187, 200 Pinard, M., 264 Pines, A., 80, 198 Pinilla, C., 82, 136 Piper, P. W., 353, 363 Pitschke, M., 167, 269 Platt, D., 268 Pocchiari, M., 17, 25, 102, 105, 166, 167, 200, 201, 257, 260, 266, 268, 269, 271, 311 Poli, G., 169, 270 Ponce, M. I., 270 Pope, A., 196 Porro, E. B., 269 Porro, M., 26, 105, 169, 197, 198, 201, 228, 270, 306 Porter, K. R., 362 Poscchiari, M., 27 Poser, S., 27, 201, 311 Post, C., 169, 270 Post, K., 143, 167, 168, 250, 269 Potempska, A., 21 Pott, U., 226, 308 Poulter, M., 305, 306, 307, 308 Power, A., 24, 80, 103, 136, 166 Poznyakovski, A. I., 332, 363 Prelli, F., 25, 26, 105, 169, 198, 200, 201, 228, 270

384

AUTHOR INDEX

Price, D. L., 224 Price, M. C., 198 Price, N. C., 23 Princen, F., 266 Priola, S. A., 1–21, 22, 24, 25, 104, 135, 136, 140, 149, 156, 159, 163, 165, 166, 167, 168, 200, 213, 227, 249, 250, 257, 261, 269, 331 Proske, D., 271 Prusiner, S., 134 Prusiner, S. B., 3, 8, 17, 21, 23, 24, 25, 26, 27, 30, 31, 33, 34, 36, 37, 39, 41, 42, 43, 45, 47, 50, 52, 53, 54, 55, 56, 57, 74, 75, 78, 79, 80, 81, 82, 83, 84, 90, 93, 96, 101, 103, 104, 105, 108, 111, 113, 124, 135, 136, 137, 139, 140, 143, 146, 148, 149, 156, 159, 161, 165, 166, 167, 168, 169, 171, 172, 180, 197, 198, 199, 200, 201, 203, 224, 225, 226, 227, 228, 231, 248, 263, 264, 265, 266, 267, 268, 269, 270, 271, 272, 285, 288, 289, 297, 304, 305, 306, 307, 308, 309, 310, 311, 327, 328, 330, 332, 338, 339, 348, 363 Ptyushkina, M. V., 361 Pucci, P., 105

Q Qin, K., 22, 79, 103, 197, 263, 304 Qin, K. F., 225 Qiu, Y., 23, 135, 228 Qu, B., 137 Quaglio, E., 199 Queitsch, C., 363

R Race, E. R., 296, 303, 309 Race, R., 14, 22, 25, 165, 264 Race, R. E., 17, 22, 25, 135, 163, 165, 167, 168, 200, 212, 225, 227 Rademacher, T. W., 45, 53 Radford, S. E., 103 Raeber, A., 21, 22, 23, 80, 103, 105, 136, 197, 198, 304, 305, 306 Raeber, A. J., 14, 17, 21, 25, 26, 192, 200, 228, 279, 304, 309 Ragg, E., 182, 200 Rahbar, F., 167 Rai, R., 317, 332 Raines, A., 168, 269

Rall, G. F., 25 Rao, C. N., 265, 269, 271 Rao, N. C., 237, 238, 239, 269 Rapp, D., 307 Raymond, G. J., 2, 4, 5, 6, 9, 15, 21, 22, 23, 24, 25, 57, 79, 80, 84, 102, 103, 104, 108, 109, 135, 136, 139–164, 165, 167, 168, 216, 219, 220, 225, 255, 258, 264, 265, 330, 331, 362 Raymond, L. D., 25, 104, 165, 168, 264 Raz, A., 268 Raz, T., 268 Reed, J. C., 267 Reekie, L. J. D., 23, 166 Regan, J. W., 269 Reid, B. G., 364 Reifenberg, K., 305 Reilly, D., 104 Reinders, L. G., 41, 43, 53 Rennard, S. I., 271 Reyes, P. F., 199, 227 Ricca, G. A., 271 Richardson, S., 105, 227 Richardson, S. L., 25, 104, 227 Richie, D., 200 Ridley, R. M., 308 Rieger, R., 236, 237, 238, 239, 240, 265, 269, 270, 271 Riek, R., 5, 6, 7, 23, 26, 27, 55–79, 80, 81, 82, 85, 86, 90, 98, 99, 102, 103, 104, 105, 110, 116, 118, 120, 126, 130, 135, 136, 137, 140, 141, 153, 155, 166, 168, 171, 184, 198, 199, 200, 201, 245, 262, 270, 280, 281, 282, 309 Riesner, D., 51, 54, 102, 104, 143, 164, 167, 168, 266, 269 Risby, D., 308 Ritchie, L. A., 12, 26, 146, 169 Rizet, G., 326, 332 Rizzardini, M., 193, 200 Robain, O., 167, 225, 265, 267, 307 Robakis, N. K., 24, 166 Robello, M., 197 Roberts, B. T., 313–329 Robertson, M. W., 267 Robey, P. G., 271 Robinson, C. V., 50, 53, 103 Robson, B., 111, 135, 136 Rocchi, M., 169, 270 Roder, D., 363 Rodgers Johnson, P., 304

AUTHOR INDEX

Rodolfo, K., 227 Rodriguez, M. L., 270 Roesler, R., 266 Rogers, M., 10, 12, 16, 23, 26, 77, 80, 135, 145, 168, 207, 224, 227, 231, 270, 306 Rohde, H., 271 Roller, P. P., 26, 81, 104, 137, 168, 200 Rosenthal, E. T., 239, 240, 270 Ross, P. D., 362 Rosseneu, M., 200 Rosseneu, M. Y., 198 Rossi, D., 306 Rossi, G., 26, 105, 169, 201, 228 Rossier, J., 167, 267, 307 Rost, B., 111, 116, 137 Rothberg, K. G., 225 Roux, B., 116, 135 Rozenshteyn, R., 82, 136 Rubenstein, R., 24, 25, 104, 146, 160, 161, 166, 167, 168, 305 Rubinstein, R., 212, 227 Rudd, P. M., 39, 53, 57, 81, 125, 137, 146, 168 Ruddock, L. W., 362 Rudelli, R. D., 224, 304 Rudiger, S., 363 Rudyk, H., 163, 168 Ruelicke, T., 270 Ruijter, J. B., 21 Rulicke, T., 23, 25, 103, 228, 309, 310 Rülicke, T., 81, 105, 271, 306 Ryan, J. B. M., 21 Rydh, A., 145, 168 Rytik, P. G., 199

S Saborio, G. P., 153, 168, 169, 200, 270 Sack, R., 50, 53 Saeki, Y., 266, 270 Safar, J., 5, 6, 8, 24, 26, 56, 59, 65, 81, 84, 104, 109, 129, 137, 141, 143, 146, 160, 161, 166, 167, 168, 169, 171, 198, 200, 201, 216, 227, 228, 269, 310 Saibil, H., 24, 103, 136, 166 Sailer, A., 21, 22, 23, 25, 80, 103, 105, 197, 200, 264, 278, 304, 305, 306, 309, 330 Saito, R., 268 Saito, S., 268 Sakaguchi, S., 204, 227, 229, 233, 270, 275, 277, 278, 308, 309, 310

385

Sakaki, Y., 135 Sakamoto, N., 227 Salas, P. J., 238, 239, 241, 270 Salès, N., 209, 227 Salmona, M., 23, 169, 171–196, 197, 198, 200, 201, 228, 270, 306 Salvatore, M., 102, 167 Sambrook, J., 245, 265 Samsonova, M. G., 361 Sanchez, H., 23, 135, 306 Sanchez, Y., 353, 354, 358, 359, 363 Sanchezsalazar, J., 228 Sander, C., 269 Sander, R., 111, 116, 137 Sandmeier, E., 50, 53, 198, 267, 307 Sanes, J. R., 266 Sano, K., 243, 270 Santoso, A., 101, 103, 104, 351, 361, 362, 363 Sargiacomo, M., 226 Sarma, R., 201 Sasaki, H., 135 Sasson, S. B., 264 Sato, H., 309 Sato, M., 239, 240, 266, 270 Saupe, S., 330 Sawicki, G. J., 363 Scala, L. J., 224 Scalici, C. L., 227 Scallet, A. C., 200 Scarlet, A. C., 195 Schafer, O., 167, 269 Schaller, H. C., 239, 240, 266 Schaller, O., 80, 136, 198 Schatton, W. F., 199 Schatzl, H. M., 8, 11, 26, 27 Schätzl, H. M., 57, 58, 69, 70, 74, 81, 82, 84, 105, 212, 227 Schellenberg, G. D., 136 Scherer, P. E., 226 Schettini, G., 197 Schibler, M. J., 267 Schilke, B. A., 363 Schirmer, E. C., 23, 103, 354, 355, 357, 358, 362, 363 Schmidt, B., 22, 197, 225, 304 Schmitt, M. C., 269 Schmittinger, S., 269 Schneider, R., 27, 82, 105 Scholtz, J. M., 104 Schoun, J., 317, 332

386

AUTHOR INDEX

Schreiber, R. D., 266 Schreiner, R., 199 Schreuder, B. E. C., 21 Schroder, H. C., 199 Schultz-Schaeffer, W. J., 22 Schulz, S. W., 263 Schulz-Schaeffer, W., 22, 79, 80, 103, 136, 197, 198, 304 Schulzschaeffer, W., 225 Schulzschaeffer, W. J., 225 Schurmann, P., 226 Schwarz, T. F., 27, 82, 105 Scorziello, A., 197 Scott, A. C., 27 Scott, J. R., 24, 226 Scott, M., 15, 17, 21, 24, 25, 26, 27, 30, 53, 74, 77, 79, 80, 81, 104, 105, 135, 136, 165, 166, 167, 168, 169, 198, 199, 201, 224, 226, 227, 228, 266, 270, 271, 274, 278, 285, 288, 289, 290, 302, 303, 304, 307, 308, 309, 310, 311, 332 Scott, M. R., 9, 15, 17, 23, 26, 80, 103, 104, 135, 159, 168, 169, 198, 226, 269, 285, 292, 293, 294, 306, 310 Scott, M. R. D., 225 Scott, R. M., 306 Sedgewick, R. D., 29, 52 Seelig, J., 201 Seiler, C., 25, 198, 200, 227 Selfridge, J., 25, 308 Selkoe, D. J., 267 Selvaggini, C., 176, 200 Seman, M., 164, 165, 263, 265 Semancik, J. S., 185, 200 Semenov, A., 26, 201, 228, 270 Senn, H., 61, 81 Sepulveda, P., 267 Serban, A., 25, 81, 102, 104, 135, 136, 167, 168, 199, 309 Serban, D., 25, 26, 81, 166, 168, 198, 199, 201, 226, 228, 269, 270, 309, 310, 332 Serban, H., 39, 53, 81, 136, 137, 167, 168, 269, 307, 311 Serio, T. R., 335–361, 363 Sern, H., 197 Serpell, L. C., 347, 348, 352, 363, 364 Servan, A., 167 Shah, T., 305 Shah, V., 266 Shaked, G. M., 8, 26, 143, 168

Shätzl, H. M., 288, 310 Shearman, M. S., 23, 198 Sheller, J. R., 309 Sherman, F., 332 Shigematsu, K., 227, 270, 308, 309, 310 Shimatake, H., 362 Shin, R. W., 267 Shirabe, S., 227, 270, 308, 310 Shirmer, E. C., 331 Shmeeda, H., 227 Shmerling, D., 105, 233, 234, 270, 281, 283, 287, 310 Shyng, S. L., 49, 53, 209, 210, 211, 227, 231, 241, 266, 270 Shyng, S.-L., 211, 225, 227 Siddle, K. C. L., 305 Sidle, K. C., 22, 225, 228 Sidle, K. C. L., 22, 80, 81, 103, 165, 166, 169, 264, 306, 307, 311 Siebert, H., 226 Sigurdsson, E. M., 145, 168 Silei, V., 195, 200 Silverman, G. L., 226, 268, 308 Silvestrini, M. C., 100, 105 Silvia, M., 304 Sima, A. A. F., 25, 199 Simms, G., 268 Simons, K., 199, 212, 227 Sims, A. A. F., 309 Singer, M. A., 363 Singh, A. C., 319, 332 Singh, N., 100, 105, 214, 227, 228 Singh, S. K., 359, 362, 363 Sisodia, S. S., 224 Sizonencko, G. I., 361 Sklaviadis, T., 164 Sletten, K., 331, 332 Smart, E. J., 201, 228, 271 Smirnov, V. N., 310, 332, 362, 363, 364 Smiroldo, S., 197, 306 Smit, A. F., 226 Smit, A. F. A., 268, 308 Smith, C. J., 10, 26, 80, 225, 264, 305 Smith, G., 23 Smith, J. F., 267 Smith, K. L., 226 Smith, M. A., 225 Smith, P. G., 27, 201, 311 Smith, R. M., 206, 227 Smith, T. F., 137

387

AUTHOR INDEX

Smits, M., 168 Smits, M. A., 21, 25, 104, 165, 168 Snider, W. D., 226 Snow, A. D., 9, 26 Sobel, M. E., 263, 264, 265, 266, 269, 271 Somero, G. N., 248, 270 Somerville, R., 25, 308 Somerville, R. A., 9, 12, 13, 26, 93, 105, 130, 137, 146, 153, 160, 161, 162, 167, 168, 169, 199, 299, 310 Sondheimer, N., 361, 363 Sonneborn, T. M., 327, 330 Soto, C., 145, 163, 168, 169, 196, 200, 257, 261, 270 Southwood, T. R. E., 21 Sozzi, G., 266 Speed, M. A., 129, 137 Spera, S., 62, 69, 72, 81 Spillantini, M. G., 198 Spilman, P. R., 23, 135 Spraker, T. R., 167 Srinivasan, A. N., 199 St, J. T., 270 Stahl, N., 4, 5, 21, 26, 30, 31, 33, 34, 36, 39, 41, 42, 43, 45, 47, 53, 57, 81, 90, 93, 104, 105, 108, 125, 137, 165, 168, 199, 206, 216, 224, 227, 228, 269 Stansfield, I., 318, 332, 341, 362, 363 Starkey, J. R., 267 Staswick, P. E., 265 Stear, M. J., 24 Steele, G. D. J., 272 Stein, R., 227 Steinberg, M., 243, 270 Stenland, C. J., 24 Steven, A. C., 105, 332, 362, 364 Stewart, L., 135 Stewart, L. R., 198 Stewart, R. S., 218, 228 Stgeorgehyslop, P. H., 227 Stieber, A., 25, 199, 226 Stierli, B., 136, 198 Stierli, E., 80 Stimson, E., 39, 53, 206, 228 Stitzel, J. D., 363 Stockel, J., 10, 26, 104, 105, 137, 169 Stöckel, J., 178, 200, 205, 228, 310 Stome, R., 103 Stone, D. E., 364 Stott, K., 321, 332

Stowring, L. E., 226, 307 Strasser, A., 267 Strathern, J. N., 330 Straub, K., 29, 52 Strauss, E. G., 271 Strauss, J. H., 271 Streit, P., 80, 136, 198 Strome, R., 22, 79, 197, 225, 226, 263, 268, 304, 308 Stubblebine, W. H., 307 Studier, F. W., 61, 81 Stultz, C. M., 137 Suarato, A., 169, 263, 270 Sugimoto, K., 310 Sugimoto, T., 227, 270, 308 Suhr, O., 168 Sulima, M. P., 304 Sunde, M., 103, 347, 363, 364 Supattapone, S., 145, 169, 175, 201, 282, 310 Surewicz, W. K., 104, 105, 136, 137, 167, 169 Surguchov, A. P., 362 Suttie, A., 22, 165, 305 Suttle, A., 197 Suzuki, H., 268 Suzuki, S., 270 Swietnicki, W., 86, 87, 91, 92, 97, 98, 101, 104, 105, 109, 126, 134, 136, 137, 148, 167, 169 Sy, M. S., 25 Sy, M.-S., 308 Sykes, B. D., 72, 82 Sznchez, H., 135 Szyperski, T., 81

T Tagliabue, E., 263, 264 Tagliavini, F., 17, 18, 20, 23, 25, 26, 65, 67, 74, 100, 105, 145, 163, 166, 169, 171–196, 197, 198, 199, 200, 201, 208, 222, 227, 228, 257, 261, 270, 306 Tait, L., 268 Takayama, S., 267 Takeshita, S., 269 Taketani, S., 270 Takeuchi, H., 25, 199, 226 Talussot, C., 200 Tanaka, K., 270 Tanaka, S., 267

388

AUTHOR INDEX

Tang, Z., 226 Tanihara, H., 270 Taraboletti, G., 264 Taraboulos, A., 4, 6, 10, 12, 21, 25, 26, 103, 135, 137, 165, 168, 185, 198, 199, 201, 208, 212, 216, 219, 220, 221, 222, 224, 225, 226, 227, 228, 260, 264, 270, 271 Taraboulos, S., 198 Tarentino, A., 30, 53, 225 Tarunina, O. V., 361 Tateishi, J., 24, 25, 104, 135, 167, 198, 267, 286 Tatzelt, J., 21, 26, 153, 163, 169, 227, 244, 245, 246, 248, 252, 263, 271, 310 Taulien, J., 363 Taylor, B. A., 305 Taylor, K. I., 347, 364 Taylor, K. L., 105, 313–329, 332 Taylor, L., 26, 81, 105, 310 Telckov, M. V., 362 Teller, J. K., 22, 225, 264 Telling, G., 26, 103, 224 Telling, G. C., 8, 9, 15, 20, 23, 25, 26, 27, 75, 77, 81, 98, 105, 110, 124, 125, 137, 160, 161, 162, 169, 232, 241, 245, 247, 271, 290, 291, 292, 298, 308, 310 Tempst, P., 30, 52, 81, 167, 269 Teplow, D., 30, 33, 36, 39, 53, 225 Teplow, D. B., 22, 27, 30, 52, 53, 81, 82, 105, 137, 167, 169, 225, 228, 264, 269 Ter Avanesyan, M. D., 300, 310 Ter-Avanesyan, M. D., 332, 338, 343, 344, 345, 357, 358, 361, 362, 363, 364 Terada, K., 268 TerAvanesyan, A., 319, 332 Terranova, V. P., 269 Terwilliger, J. D., 307 Terzi, E., 193, 201 Thellung, S., 197 Thinakaran, G., 272 Thomas, P. J., 107, 137 Thomson, V., 226, 307 Thorne, E. T., 167 Thornton, J. M., 136 Thresher, W., 22, 264 Thyer, J. M., 135 Timasheff, S. N., 248, 265 Timpl, R., 254, 271 Tings, T., 226, 307 Tinuper, P., 308

Tipler, C., 102, 224 Tisone, G. C., 168 Tissieres, A., 361 Tittmann, P., 104, 105, 331, 362 Titz, S., 306 Tjerberg, L. O., 331 Tobler, I., 57, 81, 205, 228, 229, 271, 275, 311 Tokuda, T., 199 Tome, F., 263 Tomokane, N., 267 Tomoyasu, T., 363 Torchia, M., 23, 24, 25, 26, 27, 80, 81, 103, 135, 166, 168, 169, 198, 201, 226, 271, 306, 307, 308, 309, 310, 311, 332 Torchia, T., 270 Torhia, M., 105 Traber, R., 81 Tranchant, C., 199 Traub, R. D., 185, 201 Tremblay, P., 23, 24, 26, 27, 80, 135, 166, 198, 226, 267, 268, 280, 306, 307, 308, 310 Trembly, P., 103 Tritschler, H. J., 308 Trittmann, P., 80 Trojanowski, J. Q., 25, 199, 309 Trus, B. L., 362 Tsujimura, A., 269 Tu, Y.-H., 226 Tuite, M. F., 319, 330, 332, 336, 359, 361, 362, 363, 364 Turcq, B., 325, 332 Turk, E., 9, 27, 30, 53, 57, 82, 93, 105, 130, 137, 154, 169, 206, 228 Turner, A. J., 225 Turoscy, V., 317, 332 Turq, B., 330 Tuzi, N. L., 25, 308 Tyler, A. N. J., 29, 52 Tzagoloff, A., 265

U Udy, H. J., 21 Ulyanov, N. B., 24, 80, 104, 136, 167 Uratani, Y., 348, 364 Urig, C. B., 25, 104, 225, 227 Ushijima, H., 199, 269

AUTHOR INDEX

V van Dam, A. M., 201 van den Brule, F., 264 van Dijken, P., 49, 53, 225 van, D. R., 267 Van, N. W., 266 Vancleff, J., 228 Vandekerckhove, J., 200 Vandlen, R., 104 Vanlao, B., 200 Varasi, M., 169, 270 Vasiljevic, S., 168 Veiga, S. S., 266, 268 Venturini, G., 200 Verga, L., 166, 197, 198, 201, 306 Vergnes, J. P., 264 Verkman, A. S., 263 Vey, M., 24, 185, 201, 211, 221, 226, 228, 231, 232, 241, 266, 271 Vidal, R., 198 Vidugiriene, J., 226 Viles, J. H., 23, 80, 82, 96, 102, 103, 105, 135, 166, 179, 197, 201, 205, 228, 265, 306 Villare, F., 308 Vinters, H. V., 200, 227 Vital, C., 22, 103 Vnencak-Jones, C. L., 22, 103, 309 Voellmy, R., 271 Vogel, J. L., 363 Vogel, M., 22, 225 Voigt, S., 226 Voigtlander, T., 21, 304 von Bohlen, A., 79, 103, 197, 304 von Bohlens, A., 22 von Brunn, A., 27, 82, 105 von der Mark, K., 267 von Mering, C., 270 von Schroetter, C., 27, 80, 82, 105, 137, 201 von Schrötter, C., 80 von, B. A., 263 Vonbohlen, A., 225

W Wade, W. F., 199 Wadsworth, J. D., 102, 105, 161, 169 Wadsworth, J. D. F., 180, 201 Waggoner, 205

389

Wagner, G., 63, 65, 81, 82 Wagner, J. S., 168 Walchli, M., 26, 167, 304, 310 Wälchli, M., 30, 52, 53, 79, 81, 135 Walencewicz, A. J., 200 Walker, C. A., 9, 24, 162, 163, 167, 255, 256, 266 Wallace, A. C., 26, 80, 136, 166, 167, 200, 228, 266, 269 Wallace, A. L., 104 Walter, J., 304 Walter, W. A., 363 Walther, D., 26, 310 Waltho, J. P., 24, 80, 103, 136, 166 Walz, R., 266 Wandosell, F., 199 Wang, D. I. C., 137 Wang, J. L., 267 Wang, K., 226, 268, 308 Wang, K. S., 239, 240, 271 Wang, R., 47, 49, 53, 225 Wang, S., 104 Wang, Y. H., 271 Warter, J. M., 199 Warwicker, J., 124, 137, 250, 271 Wataya, K. M., 266 Watt, C. J., 21 Weaver, D., 165 Weber, C., 81 Weber, D. J., 50, 53 Weber-Ban, E. U., 355, 364 Weeks, B. S., 266 Wegrzyn, R. D., 363 Wehlburg, C. M., 168 Wehrly, K., 22, 135, 165, 269 Weidenhofer, G., 27, 105 Weidenhöfer, G., 82 Weiner, R. I., 26, 227 Weinman, N. A., 268 Weiss, S., 9, 229–262, 265, 267, 269, 270, 271 Weissman, J. S., 103, 104, 362, 363 Weissmann, C., 21, 22, 23, 25, 30, 52, 53, 55, 74, 79, 80, 81, 82, 83, 103, 105, 135, 140, 146, 167, 169, 197, 200, 204, 225, 228, 229, 264, 270, 271, 304, 305, 306, 309, 310, 330 Welch, W. J., 169, 221, 228, 248, 263, 271 Wells, G. A. H., 1, 21, 27 Wemmer, D. E., 80, 198

390

AUTHOR INDEX

Wen, G. Y., 224 Wendler, W., 265 Werner, B., 81 Werner, T., 27, 82, 105 Werner-Washburne, M., 354, 358, 359, 363, 364 West, J. D., 226, 307 Westaway, D., 15, 22, 25, 26, 27, 30, 52, 53, 79, 81, 103, 135, 167, 168, 197, 225, 226, 263, 268, 287, 296, 302, 303, 304, 305, 307, 308, 309, 310, 311, 328, 332 Westermark, G. T., 331 Westermark, P., 328, 329, 331, 332 Wewer, U., 266 Wewer, U. M., 240, 269, 271 Wheeler, S. F., 39, 53, 81, 137, 168 White, A. R., 135, 198 White, J. V., 116, 137 Whittal, R. M., 50, 54, 102, 105 Whittington, M. A., 80, 205, 225, 228, 264, 278, 305, 307, 311 Wicha, M. S., 237, 239, 267 Wickner, R. B., 8, 27, 105, 300, 311, 313–329, 331, 332, 333, 337, 338, 339, 352, 362, 364 Wickner, S., 364 Wider, G., 26, 27, 79, 80, 81, 82, 102, 103, 104, 105, 135, 137, 166, 168, 200, 201, 262, 270, 309 Wiedmann, M., 363 Wieland, F., 199 Wildegger, G., 90, 103, 105 Wilder, G., 80 Wilesmith, J. W., 1, 21 Will, H., 201 Will, R., 2, 25, 308, 310 Will, R. G., 22, 25, 27, 104, 165, 168, 173, 197, 201, 305, 311 Wille, H., 24, 81, 167, 168, 169, 198, 201, 228, 269, 271 Willett, W. S., 104 Williams, A., 200 Williams, A. E., 188, 194, 201 Williams, A. F., 34, 53 Williams, E. S., 2, 27, 167, 168 Williams, R. W., 105, 332, 364 Williamson, R. A., 77, 82, 136 Willie, H., 166, 310 Willmer, J., 26 Wilson, P. G., 342, 344, 364

Windl, O., 226 Winklhofer, K. F., 163, 169 Winnacker, E. L., 265, 271 Winnacker, E.-L., 271 Wirth, P., 271 Wishart, D. S., 62, 72, 82 Wisniewski, H., 136 Wisniewski, H. M., 24, 166, 167, 198, 199 Wisniewski, T., 168, 169, 200, 270 Wong, B.-S., 22, 264 Wong, C., 139–164 Wong, C. W., 253, 265 Wong, J. M., 272 Wong, K., 212, 228 Wong, S. F., 29, 52 Wong, S. K., 29, 52 Wood, S. C., 26, 166, 169 Wood, S. C. E. R., 310 Woolhouse, M. E. J., 21 Wopfner, F., 14, 27, 57, 69, 70, 74, 82, 105 Wordeman, L., 239, 240, 270 Wormald, M. R., 39, 53, 81, 137, 168 Wrenn, D. S., 266 Wright, P. E., 23, 80, 82, 103, 105, 135, 166, 197, 201, 228, 265, 306 Wustemann, F. S., 263 Wuthrich, K., 23, 26, 27, 103, 105, 135, 136, 137, 262, 270, 309, 331, 362 Wüthrich, K., 55–79, 80, 81, 82, 102, 104, 137, 140, 141, 153, 166, 168, 198, 199, 200, 201

X Xi, Y. G., 167, 257, 260, 268, 271 Xia, T., 79 Xiong, L.-W., 139–164 Xue, R., 308

Y Yamaguchi, K., 24, 104 Yanai, A., 137, 227, 264 Yang, E., 253, 271 Yang, R. Y., 237, 272 Yang, S., 81 Yang, S. L., 25, 26, 168, 271, 307, 309, 311 Yang, S.-L., 309, 310, 332 Yansura, D., 104 Yao, J., 82

AUTHOR INDEX

Ye, X., 195, 200, 305 Yehiely, F., 23, 26, 135, 168, 236, 252, 272, 306 Yin Qiu, 306 Ying, Y.-S., 225 Yokoyama, T., 265 Yost, C. S., 24, 218, 225, 226, 228, 284, 311 Young, C., 338, 344, 364 Young, K., 25, 174, 177, 198, 199, 200, 201, 227, 309 Young, S., 2, 27 Yow, H., 237, 239, 272

Z Zahn, R., 6, 16, 18, 23, 27, 61, 71, 72, 73, 80, 82, 84, 85, 99, 105, 110, 120, 127, 137, 171, 199, 201 Zaidi, S. I., 225 Zaitsev, I., 196, 268

391

Zanata, S. M., 266, 268 Zanusso, G., 105, 214, 227, 228 Zebarjadian, Y., 311 Zeidler, M., 27, 201, 311 Zhang, H., 24, 65, 80, 82, 87, 91, 93, 104, 105, 109, 113, 120, 136, 137, 148, 166, 169, 183, 184, 198, 201 Zhao, D., 136 Zhouravleva, G., 318, 333, 364 Ziegelhoffer, T., 361, 363 Zimmerman, T. R., 200, 227 Zink, A. D., 361 Zobeley, E., 87, 94, 95, 101, 105 Zolkiewski, M., 352, 355, 364 Zorzoli, I., 268 Zou, W., 22, 103, 199, 227 Zuegg, J., 128, 137 Zuiderweg, E. R. P., 62, 80 Zulianello, L., 16, 27, 80, 104, 136, 166, 266 Zuo, J., 271

316

REED B. WICKNER ET AL.

TABLE II Phenotypic Relation of a Prion or a Nucleic Acid Replicon with Chromosomal Genesa Nonmendelian Element

Presence of Nonmendelian Elementa

Chromosomal Mutant that Loses the Elementb

Relation of a and b

Does Replacing the Mutant Gene Restore the Phenotype?

opposite opposite same same same same

no no no yes yes yes

Phenotypes M dsRNA mitDNA mitDNA-DI Prion [URE3] [PSI]

killer + glycerol + glycerol – defective USA uptake + suppressor⇑

killer – glycerol – glycerol – defective USA uptake + suppressor⇑

a M dsRNA is a dsRNA virus of yeast encoding a secreted toxin (the killer toxin). The mitochondrial DNA (mitDNA) encodes proteins needed for oxidative phosphorylation and thus for growth on glycerol as a carbon source. Some deletion mutations of mitDNA are defective for function but interfere with the replication of the normal mitDNA and so make their presence known. Replacing a chromosomal gene needed for replication of mitDNA or M dsRNA or the defective interfering mitDNA-DI does not restore the killer or glycerol phenotype to “+” because the nucleic acid replicon is still missing. But a prion producing a phenotype because of the absence of the normal form has the same phenotype as mutation of the gene for the protein. Replacing the gene for the protein replaces the normal form and so the phenotype is restored. These properties allow the geneticist to distinguish nucleic acid replicons from prions (Wickner, 1994).

that both were prions (Wickner, 1994). We also detail the more recent studies of [URE3] and the new prion, [Het-s]. More information concerning [PSI] is provided in the chapter by Serio and Lindquist et al., this volume. III. [URE3] AND URE2 AFFECT NITROGEN CATABOLITE REPRESSION Yeast cells presented with a rich nitrogen source, such as ammonia or glutamine, shut off production of enzymes necessary for the utilization of poor nitrogen sources, such as allantoate or urea. A poor nitrogen source, such as proline, does not shut off utilization of other poor sources. This phenomenon is called “nitrogen catabolite repression” or “nitrogen regulation” (Cooper, 1982; Magasanik, 1992). Lacroute wanted to supplement mutants blocked in the first step of uracil biosynthesis, aspartate transcarbamylase (ura2), with the product of the reaction, ureidosuccinate (USA). He found that when ammonia was supplied as the nitrogen source, USA was not taken up, but that on

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SUBJECT INDEX

A

URE2, nitrogen catabolite repression, 316–318 URE3 nitrogen catabolite repression, 316–318 prion evidence, 321–322, 329 URE2p relationship, 319, 322–323 URE2p amyloid formation in vitro, 323–325 prion domains, 319–321 protease resistance, 322–323 URE3 relationship, 319, 322–323, 329 amyloid proteins in fatal familial insomnia, 173 formation, in vitro, 323–325 in Gerstmann-Sträussler-Schienker syndrome, 172–175, 188–194 anthracycline 4′-iodo-4′-deoxy-doxorubicin, anti-TSE propertiesanti-TSE properties, 257, 259, 261 astroglial cultures, transmissible spongiform encephalopathy pathogenesis, prion peptide fragment studies, 194–196

aging, prion protein modification analysis, 50 α-helical proteins computational analysis, 113–120 conformational marker, 75–76, 109, 175 conversion to β-sheet, 92–93, 109, 141, 175 globular domain structure, 68, 71, 85, 171 infrared spectra, 59–60 strain differences, 109–113, 154, 175–185 amphotericin B, anti-TSE properties, 163, 257–260 amyloid diseases, see also transmissible spongiform encephalopathies; specific diseases yeast prion models, 313–329 genetic criteria, 314–316 gene mutation link to phenotypes, 315–316 overproduction link to prion emergence, 315 reversible curability, 314–315 het-s, 325–327, 329 nitrogen catabolite repression, 316–318 overview, 313–314, 329 Podospora anserina prions amyloidoses implications, 328–329 function, 325–327 PSI SUP35p relationship, 319 translation termination efficiency, 318 SUP35, translation termination efficiency, 318 SUP35p, PSI relationship, 319 translation termination efficiency, 318 TSE prions compared, 327–328

B Bax protein, prion protein interactions, 253–254 Bcl-2 protein, prion protein interactions, 253–254 β-sheet in prion proteinC conversion from α-helical protein, 92–93, 109, 141, 175 globular domain structure, 68–71, 85, 171 infrared spectra, 59–60 in prion protein-res, 2, 141 in prion proteinSc 393

394

SUBJECT INDEX

β-sheet (continued) elevated content, 56, 109, 113, 171 stability, 109 in prion protein-sen, 154 strain differences, 109–113, 175–185 biosynthesis overview, 4–6, 203–204, 222 prion proteinC, 204–212 clathrin-coated pits, caveolae and rafts, 210–212, 231 expression and function, 204–206 mechanisms, 206–208 posttranslational cleavage, 208 structure, 206–208 subcellular localization and trafficking, 208–210 topological forms, 284–286 prion protein-res, 4–6 prion proteinSc cell culture models, 212–215 membrane attachment mechanisms, 216–218, 231–235 posttranslational cleavage, 222 production kinetics, 141, 218–220 subcellular formation site, 220–222 subcellular localization, 215–216 prion protein-sen, 4 blood-brain barrier, anti-TSE drug research, 163 bovine prion protein globular domain three-dimensional structure, 62–65, 73, 77 transgenic species barrier studies bovine to human transmission, 294–295 bovine to murine transmission, 292–294 bovine spongiform encephalopathy Creutzfeldt-Jakob disease link, 2, 173, 293–295 description, 1–2, 273–274 prion protein role, 15, 301 protein-only hypothesis, 83, 96, 248, 275 C cadherins, prion protein interactions, 242–243 carrageenan, anti-TSE properties, 255 cell membranes

prion proteinSc biosynthesis, attachment mechanisms, 216–218, 231–235 transmissible spongiform encephalopathy pathogenesis prion peptide fragment studies, 185–187 surface receptors role, 231–243 cadherins, 242–243 37 kDa laminin receptor precursor, 230, 237–242 66 kDa membrane protein, 235–236 receptor function, 231–235 chaperone molecules prion protein interactions, 243–248 chemical chaperones, 248 heat shock proteins, 245–247, 353–359 protein X, 247–248 PSI metabolism regulation, 352–360 Hsp70, 354 Hsp104, 353–359 inheritance role, 356–357 maintenance role, 356–357 models, 357–358 Ssa1, 358–359 Ssb, 359–360 Chinese hamster ovary cells, prion proteinSc biosynthesis, 212–215, 219, 221 chronic wasting disease description, 2 prion protein role codon 129 homozygosity, 15 PrPC conversions, 157 circadian rhythms, prion protein role, 57, 229 circular dichroism spectroscopy, prion protein, 57-59,84-89,109,178 clathrin-coated pits and vesicles, endocytic prion protein recycling pathway, 210–212, 231 clusterin, anti-TSE properties, 260–261 conformations molecular chaperones role, 243–248 chemical chaperones, 248 heat shock proteins, 245–247 protein X, 247–248 prion peptide fragment analysis PrPC to PrPSc conversion study, 175–185 globular domain, 182–185

395

SUBJECT INDEX

N-terminal domain, 171, 173, 177–182 octapeptide repeat region, 177–180 simulations and computational analysis, 110–118 recombinant prion protein folding dynamics, 83–102 disulfide bond role, 93–96 inherited human prion diseases, 96–101 mechanisms, 85–93 overview, 83–85, 101–102 point mutations, 96–101 thermodynamic stability, 96–101 two-state model, 86–87, 90–91 unfolding studies, 141–145 simulations and computational analysis of prion proteins, 107–134 atomic-level molecular dynamics simulations, 128–134 human disease causing mutations, 125–128 nuclear magnetic resonance imaging, 109, 118–134, 180, 184 peptide fragment studies, 110–118 PrPC to PrPSc conversion, 109, 129–134 species barriers, 123–124 static modeling, 121–128 strain differences, 124–125 structure prediction models, 110–112, 116, 118–121 transitions, 107–110 Congo red, anti-TSE properties, 163, 257–259 copper II mass spectrometric analysis, 50–51, 57 prion protein binding mechanisms, 10, 179–180, 205, 224 transport role, 177–180 Cp-60 and -62, anti-TSE properties, 257, 261 Creutzfeldt-Jakob disease, see also drug research; human prion protein bovine spongiform encephalopathy link, 2, 173, 293–295 genetic link, 283–284 pathogenesis study using peptide fragments, 185, 193–194 prion protein role codon 129 homozygosity, 15 insertion mutations, 19–20

strain types, 297–300 protein-only hypothesis, 83, 96, 248, 275 C-terminal sequence, of prion protein GPI anchor interactions, 34–36, 41–42 mass spectrometric analysis, 34–36, 41–42 three-dimensional structure, 57–59, 68, 85–93 cystic fibrosis transmembrane regulator, molecular chaperone role, 248 D dapsone, anti-TSE properties, 260–261 dextran sulfate, anti-TSE properties, 163, 255–256, 258 dimethyl sulfoxide, molecular chaperone role, 248 disulfide bonds, in prion proteins, 9, 30, 93–96, 153–154 DNA, mass spectrometry analysis, 30 doppel protein, function, 232–234, 276–277 drug research, prion protein interactions, 162–163, 255–262 amphotericin B, 163, 257–260 anthracycline 4′-iodo-4′-deoxydoxorubicin, 257, 259, 261 carrageenan, 255 clusterin, 260–261 Congo red, 163, 257–259 Cp-60, 257, 261 Cp-62, 257, 261 dapsone, 260–261 dextran sulfate, 163, 255–256, 258 flurpirtin, 260–261 heparin, 230, 255 heteropolyanion (23), 163, 255–256 iPrP13, 257, 261 MS-8209, 257–260 overview, 162–163, 230–231 pentosan polysulfate, 162, 255–256, 258–259 phtalocyans, 257, 261 polyanions, 255–258 polyene antibiotics, 258–260 porphyrins, 163, 257, 261 pythalocyanines, 163 E electrospray ionization mass spectrometry, prion protein analysis, 29–52

396

SUBJECT INDEX

electrospray ionization mass spectrometry, prion protein analysis (continued) accessory molecules in scrapie, 51 aging effects, 50 copper binding, 50–51, 57 C-terminal analysis, 34–36, 41–42 GPI anchor analysis, 39–45 C-terminal peptide-GPI analysis, 34–36, 41–42 glycan branching pattern analysis, 42–43 unanswered questions, 43–45, 52 identification, 30–31 intact proteins, 45–49 mammal and chicken PrP compared, 49 N-linked oligosaccharides, 30, 37–39, 46 non-PrP peptides in preparations, 37 N-terminal analysis, 29, 31, 33–35, 49 overview, 29–30, 51–52 preliminary analysis, 30–31 PrPSc amino acid sequence confirmation, 31–37 recombinant proteins, 49–51 synthetic peptides, 49–51 endocytic recycling pathway clathrin-coated pits and vesicles role, 210–212, 231 mechanisms, 209–212

bovine protein, 62–65, 73, 77 human protein, 71–73, 76–78 murine protein, 65, 68–71, 77, 87 Syrian hamster protein, 71, 74, 176 glutamate neurotoxicity, 195 PSI metabolism, 345 glycerol, molecular chaperone role, 248 glycosaminoglycans endocytic prion targeting, 211 PrP-sen conversion to PrP-res, 9 glycosylation, in prion protein synthesis, 10, 12–13, 207–208 glycosyl phosphotidylinositol anchor mass spectrometric prion protein analysis, 39–45 C-terminal peptide-GPI analysis, 34–36, 41–42 glycan branching pattern analysis, 42–43 protein identification, 30 unanswered questions, 43–45, 52 phospholipase interactions, 30–31, 186, 209 in prion protein synthesis, 206–210 three-dimensional structure, 57–59 in transmissible spongiform encephalopathies, 4, 10, 186

F

H

fast atom bombardment mass spectrometry, prion protein analysis, 29, 32 fatal familial insomnia, amyloid protein role, 173 flurpirtin, anti-TSE properties, 260–261 folding dynamics, see conformations Fourier transform infrared spectroscopy, prion protein three-dimensional structure, 109–110, 182–185

hamster studies, see Chinese hamster ovary cells; Syrian hamster prion protein heat shock proteins molecular chaperone role, 245–247 PSI metabolism regulation, 353–359 heparin, anti-TSE properties, 230, 255 heteropolyanion (23), anti-TSE properties, 163, 255–256 het-s, function, 325–327, 329 1H-1H nuclear Overhauser effect spectroscopy, prion protein, 62-65 high-performance liquid chromatography, prion protein analysis, 31–37 human prion protein, see also CreutzfeldtJakob disease description, 274 globular domain three-dimensional structure, 71–73, 76–78 pathogenic mutations, 283–284 transgenic studies

G Gerstmann-Sträussler-Schienker syndrome amyloid peptide role, 172–175, 188–194 prion protein role, 19–20, 213–214, 222 globular domains, of prion protein PrPC to PrPSc conversion study, 182–185 three-dimensional structure α-helical proteins, 68, 71, 85, 171 β-sheets, 68–71, 85, 171

397

SUBJECT INDEX

conditional knockouts, 279–280 disease link, 283–284 species barrier studies bovine to human transmission, 294–295 human to murine transmission, 290–292

I infrared spectroscopy, prion protein threedimensional structure, 59–60, 109–110, 182–185 iPrP13, anti-TSE properties, 257, 261

K knockout genes, see transgenics

L laminin 37 kDa laminin receptor precursor, 230, 237–242 prion protein interactions, 230, 237–242, 254–255 ligands, of prion protein, 251–253 (Pli 3-8), 252–253 (Pli 45), 251–252 (Pli 110), 251–252 liquid secondary ionization mass spectrometry, prion protein analysis, 29, 32–35

M mad cow disease, see bovine spongiform encephalopathy manganese II, prion protein function role, 10 mass spectrometry collision-induced dissociation, 30, 42–43 prion protein analysis, 29–52 accessory molecules in scrapie, 51 aging effects, 50 copper binding, 50–51, 57 C-terminal analysis, 34–36, 41–42 GPI anchor analysis, 39–45 C-terminal peptide-GPI analysis, 34–36, 41–42 glycan branching pattern analysis, 42–43

unanswered questions, 43–45, 52 identification, 30–31 intact proteins, 45–49 mammal and chicken PrP compared, 49 N-linked oligosaccharides, 30, 37–39, 46 non-PrP peptides in preparations, 37 N-terminal analysis, 29, 31, 33–35, 49 overview, 29–30, 51–52 preliminary analysis, 30–31 PrPSc amino acid sequence confirmation, 31–37 recombinant proteins, 49–51 synthetic peptides, 49–51 matrix-assisted laser desorption/ionization, prion protein analysis, 29, 45–51 membranes, see cell membranes metals mass spectrometric analysis, 50–51, 57 prion protein binding mechanisms, 10, 179–180, 205, 224 transport role, 177–180 microglial cultures, transmissible spongiform encephalopathy pathogenesis, prion peptide fragment studies, 194–196 molecular chaperones, see chaperone molecules MS-8209, anti-TSE properties, 257–260 murine prion protein globular domain three-dimensional structure, 65, 68–71, 77, 87 production in null mice neurografting experiments, 278–279 transgenic species barrier studies bovine to murine transmission, 292–294 human to murine transmission, 290–292 murine to hamster transmission, 288–289

N neurografting experiments, PrP-producing cells in null mice, 278–279 neuronal culture analysis, transmissible spongiform encephalopathy pathogenesis, prion peptide fragment studies, 187–194

398

SUBJECT INDEX

neurotoxicity, of prion proteins, 20, 146, 187–195, 300–302 N-glycosidase F, prion protein identification, 30, 37–39, 46 nitrogen catabolism, repression by yeast prions, 316–318 N-linked oligosaccharides mass spectrometric prion protein analysis, 30, 37–39, 46 in prion protein synthesis, 206–207 three-dimensional structure, 57 N-terminal sequence, of prion protein flexible tail, 16, 73–74, 180–182 mass spectrometric analysis, 29, 31, 33–35, 49 octapeptide repeat region, 177–180, 206 PrPC to PrPSc conversion study, 171, 173, 177–182 PSI inheritance in yeast, 345 three-dimensional structure, 57–59, 69, 85–93 transgenic studies, 280–281 nuclear magnetic resonance imaging, of prion proteins conformation analysis overview, 109–110 static models, 121–134 atomic-level dynamics, 128–134 features, 121–123 human disease-causing mutations, 125–128 internal deletions, 281–283 PrPC to PrPSc conversion simulation, 109, 118–134, 180, 184 PrPC two molecular dynamic simulations, 128–129 species barriers, 123–124 strain differences, 124–125 structural models, 110–112, 116, 118–121 three-dimensional analysis, 56, 59–74, 85–86 nucleic acids mass spectrometry analysis, 30 prion protein interactions, 262

O octapeptide repeat region of prion peptide fragments conformation analysis, PrPC to PrPSc conversion study, 177–180

in prion protein synthesis, 206, 283–284 oligosaccharides, N-linked mass spectrometric prion protein analysis, 30, 37–39, 46 in prion protein synthesis, 206–207 three-dimensional structure, 57

P pentosan polysulfate, anti-TSE properties, 162, 255–256, 258–259 peptide fragments, of prion proteins amyloid peptides in GerstmannSträussler-Schienker syndrome, 172–175, 188–194 conformation analysis PrPC to PrPSc conversion study, 175–185 globular domain, 182–185 N-terminal domain, 171, 173, 177–182 octapeptide repeat region, 177–180 simulations and computational analysis, 110–118 overview, 171–173, 196 pathogenesis study, 185–196 cell membrane interactions, 185–187 neuronal culture analysis, 187–194 astroglial cultures, 194–196 human disease-associated mutations, 193–194 microglial cultures, 194–196 (PrP 106-126), 187–193 pharmaceuticals, see drug research phosphatidylinositol phospholipase C GPI binding, 30–31, 186, 209 prion protein releasable, 213, 216 prion protein resistance, 213, 216–221, 224 phtalocyans, anti-TSE properties, 257, 261 Podospora anserina prions amyloidoses implications, 328–329 function, 325–327 polyanions, anti-TSE properties, 255–258 polyene antibiotics, anti-TSE properties, 258–260 porphyrins, anti-TSE properties, 163, 257, 261 prion proteinC, see also transmissible spongiform encephalopathies Bcl-2 interactions, 253–254

SUBJECT INDEX

biosynthesis cellular biology clathrin-coated pits, caveolae and rafts, 210–212, 231 expression and function, 204–206 life cycle, 241–242 mechanisms, 206–208 posttranslational cleavage, 208 structure, 206–208 subcellular localization and trafficking, 208–210 overview, 4–6, 203–204, 222, 242 cell surface receptor interactions, 231–243 cadherins, 242–243 37 kDa laminin receptor precursor, 230, 237–242 66 kDa membrane protein, 235–236 receptor function, 231–235 conformations molecular chaperones role, 243–248 chemical chaperones, 248 heat shock proteins, 245–247 protein X, 247–248 simulations and computational analysis, 107–134 atomic-level molecular dynamics simulations, 128–134 human disease causing mutations, 125–128 nuclear magnetic resonance imaging, 109, 118–134, 180, 184 peptide fragment studies, 110–118 PrPC to PrPSc conversion, 109, 129–134 species barriers, 123–124 static modeling, 121–128 strain differences, 124–125 structure prediction models, 110–112, 116, 118–121 transitions, 107–110 conversions, 139–164 binding versus conversion of heterologous molecules, 157–159 in chronic wasting disease, 157 disulfide bond role, 153–154 efficiency versus susceptibility, 156–157 isoform interactions, 248–251 isoform nomenclature, 140–141 overview, 139–141, 164

399

perturbations, 162 PrPC to PrPSc conversion simulations and computational analysis, 109, 129–134 species barriers, 13–17, 156–159 in transmissible spongiform encephalopathies anti-TSE drug research, 162–163 deletion forms, 145 disease strains, 159–162 infectivity, 162 mechanisms, 139–141 structural diversity, 145–146 C-terminal sequence, three-dimensional structure, 57–59, 68, 85–93 disease mechanisms, 2–3, 19–20 disulfide bonds, 30, 84, 93–96, 153–154 drug interactions, 162–163, 255–262 amphotericin B, 163, 257–260 anthracycline 4′-iodo-4′-deoxydoxorubicin, 257, 259, 261 carrageenan, 255 clusterin, 260–261 Congo red, 163, 257–259 Cp-60, 257, 261 Cp-62, 257, 261 dapsone, 260–261 dextran sulfate, 163, 255–256, 258 flurpirtin, 260–261 heparin, 230, 255 heteropolyanion (23), 163, 255–256 iPrP13, 257, 261 MS-8209, 257–260 overview, 162–163, 230–231 pentosan polysulfate, 162, 255–256, 258–259 phtalocyans, 257, 261 polyanions, 255–258 polyene antibiotics, 258–260 porphyrins, 163, 257, 261 pythalocyanines, 163 familial infection mechanisms, 19–20 mutant properties, 19 glycosylation, 10, 12–13, 207–208 hosts species, 1–2 species barriers, 13–17, 156–159 virus pathogenesis, 13 isoform interactions, 248–251 isoform nomenclature, 140–141

400

SUBJECT INDEX

prion proteinC (continued) laminin interactions, 230, 237–242, 254–255 ligands, 251–253 (Pli 3-8), 252–253 (Pli 45), 251–252 (Pli 110), 251–252 neurotoxicity, 20, 146, 187–195, 300–302 N-terminal sequence mass spectrometric analysis, 29, 31, 33–35, 49 three-dimensional structure, 57–59, 69, 85–93 transgenic studies, 280–281 nucleic acid interactions, 262 overview, 1–3, 20–21, 164 peptide fragment studies, see peptide fragments recombinant proteins folding dynamics, 83–102 disulfide bond role, 93–96 inherited human prion diseases, 96–101 mechanisms, 85–93 overview, 83–85, 101–102 point mutations, 96–101 thermodynamic stability, 96–101 two-state model, 86–87, 90–91 mass spectrometric analysis, 49–51 three-dimensional structure, 57–59, 67–74 strain differences conformation analysis, 124–125 conversion analysis, 159–162 molecular basis, 297–302 description, 297 neurodegeneration mechanisms, 300–302 strain types, 297–300 structure analysis flexible tail, 73–74, 180–182 globular domain bovine protein, 62–65, 73, 77 human protein, 71–73, 76–78 murine protein, 65, 68–71, 77, 87 Syrian hamster protein, 71, 74, 176 mass spectrometric analysis, 29–52 accessory molecules in scrapie, 51 aging effects, 50 amino acid sequence confirmation, 31–37

copper binding, 50–51, 57 C-terminal analysis, 34–36, 41–42 glycan branching pattern analysis, 42–43 GPI anchor analysis, 39–45 identification, 30–31 intact proteins, 45–49 mammal and chicken PrP compared, 49 N-linked oligosaccharides, 30, 37–39, 46 non-PrP peptides in preparations, 37 N-terminal analysis, 29, 31, 33–35, 49 overview, 29–30, 51–52 preliminary analysis, 30–31 recombinant proteins, 49–51 synthetic peptides, 49–51 unanswered questions, 43–45, 52 molecular architecture, 67–68 natural prion proteins, 57–59 nuclear magnetic resonance information, 56, 59–74, 85–86 optical spectroscopy information, 65–67 overview, 2, 7, 29, 55–56, 84 prions, 56–57 recombinant prion proteins, 57–59, 67–74, 85 species barrier, 74–78 structure calculations, 62–65 transgenic studies, 280–284 disease link, 283–284 internal deletions, 281–283 N-terminal deletions, 280–281 transgenic studies, 273–304 disease propagation conditional Prnp knockout, 275–280 neurografted PrP-producing cells in null mice, 278–279 spontaneous disease, 286–288 gene ablation effects, 275–278 gene structure and function, 280–284 disease link, 283–284 internal deletions, 281–283 N-terminal deletions, 280–281 incubation studies, 295–297 molecular basis of prion strains, 297–302 description, 297 neurodegeneration mechanisms, 300–302

SUBJECT INDEX

strain types, 297–300 overview, 273–275, 304 species barrier studies, 288–295 artificial prions, 289–290 bovine to human transmission, 294–295 bovine to murine transmission, 292–294 human to murine transmission, 290–292 mechanisms, 288 murine to hamster transmission, 288–289 topology studies, 284–286 transgene vector considerations, 302–303 prion protein-res, see also transmissible spongiform encephalopathies biosynthesis, 4–6 cofactors, 9 conformations, 11–12, 21, 142 cotranslational modifications, 4, 9–10 disulfide bond, 9 folding and unfolding, 141–145 formation β-sheet role, 2, 141 models, 6–8, 146–148 PrPC deletion forms, 145 species-specific formation, 14–15 spontaneous formation, 18–19 glycosylation, 10, 12–13 GPI anchor, 4, 10 metal binding, 10 mutant PrP-sen compared, 19 posttranslational modifications, 6, 9–10 primary sequence, 14 PrP-sen conversion biological connections, 150–152 disulfide bond role, 153–154 interaction site, 153–154 mechanisms, 6–10, 153, 164 PrP-sen interactions binding, 148–150 species barriers, 13–17, 156–159 replication, 8–9, 20 resistance, 9, 12–17, 146 structural diversity, 145–146, 164 in transmissible spongiform encephalopathies disease susceptibility, 15–17 perturbations, 162

401

species barriers, 13–17, 156–159 viral strains, 10–13, 159–162 prion proteinSc, see also scrapie; transmissible spongiform encephalopathies amino acid sequence confirmation, 31–37 biosynthesis cellular biology, 212–222 cell culture models, 212–215 membrane attachment mechanisms, 216–218, 231–235 posttranslational cleavage, 222 production kinetics, 141, 218–220 subcellular formation site, 220–222 subcellular localization, 215–216 overview, 4–6, 203–204, 222 conversions binding versus conversion of heterologous molecules, 157–159 induced conversions, 150–152 isoform interactions, 248–251 disease pathogenesis study using peptide fragments, 185–196 cell membrane interactions, 185–187 neuronal culture analysis, 187–194 astroglial cultures, 194–196 human disease-associated mutations, 193–194 microglial cultures, 194–196 (PrP 106-126), 187–193 disulfide bond, 30, 93–96, 153–154 formation, 145, 212–215 peptide fragment studies, see peptide fragments self-replication, 84–85 strain differences conformation analysis, 124–125 conversion analysis, 159–162 molecular basis, 297–302 description, 297 neurodegeneration mechanisms, 300–302 strain types, 297–300 structural analysis diversity, 124, 145–146, 164 folding dynamics β-sheet, 56, 109, 113, 171 conformation prediction, 112, 124–125

402

SUBJECT INDEX

prion proteinSc (continued) disulfide bond role, 93–96 inherited human prion diseases, 96–101 mechanisms, 85–93 overview, 83–85, 101–102 point mutations, 96–101 thermodynamic stability, 96–101 unfolding studies, 141–145 mass spectrometric analysis, 31–37, 47–48, 49–51 three-dimensional structure, 56–59, 67–74 prion protein-sen, see also transmissible spongiform encephalopathies biosynthesis, 4 conversion to PrP-res biological connections, 150–152 disulfide bond role, 153–154 interaction site, 153–154 mechanisms, 6–10, 153, 164 disulfide bond, 9, 153–154 glycosylation, 10, 12–13 mutant PrP-res compared, 19 PrP-res interactions binding, 148–150 species barriers, 13–17, 156–159 in transmissible spongiform encephalopathies disease susceptibility, 15–17 perturbations, 162 species barriers, 13–17, 156–159 viral strains, 10–13, 159–162 protease, see also prion protein-res prion protein resistance, 9, 12–17, 146 prion protein sensitive, see prion proteinsen yeast prion resistance, 322–323 proteinase K, prion protein reactions digestion, 2–3, 8, 19, 172–173 induced conversions, 151–152 unfolding studies, 141–143 protein-only hypothesis, description, 83, 96, 248, 275 protein X conversion factor role, 77 molecular chaperone role, 247–248 PrP-res formation inhibition, 156 species barrier role, 123–124 PSI description, 335–336, 360

inheritance contributions from N region, 344–345 nonapeptide repeats, 344–345 N-terminus glutamines, 345 metabolism regulation by molecular chaperones, 352–360 Hsp70, 354 Hsp104, 353–359 inheritance role, 356–357 maintenance role, 356–357 models, 357–358 Ssa1, 358–359 Ssb, 359–360 modeling in vitro, 346–352 alternate protein state replication, 347–350 N mutation biochemical effects, 350–352 SUP35 self-assembly, 346–347 phenotype model, 341–342 prion evidence, 338–341 alternative phenotype associations, 339 alternative physical state replication, 339–341 SUP35 link, 338–339 SUP35 overexpression induction, 339 prion hypothesis formulation, 336–338 SUP35p relationship, 319 translation termination efficiency, 318 pythalocyanines, anti-TSE properties, 163

R RNA, mass spectrometry analysis, 30

S Saccharomyces cerevisiae, see yeast prions scrapie, see also prion proteinSc description, 2 protein-only hypothesis, 83, 96, 248, 275 resistance, 9, 12–17 therapeutic drugs, 162–163, 255–262 amphotericin B, 163, 257–260 anthracycline 4′-iodo-4′-deoxydoxorubicin, 257, 259, 261 carrageenan, 255 clusterin, 260–261 Congo red, 163, 257–259 Cp-60, 257, 261 Cp-62, 257, 261

403

SUBJECT INDEX

dapsone, 260–261 dextran sulfate, 163, 255–256, 258 flurpirtin, 260–261 heparin, 230, 255 heteropolyanion (23), 163, 255–256 iPrP13, 257, 261 MS-8209, 257–260 overview, 230–231 pentosan polysulfate, 162, 255–256, 258–259 phtalocyans, 257, 261 polyanions, 255–258 polyene antibiotics, 258–260 porphyrins, 163, 257, 261 pythalocyanines, 163 transgenic studies, see transgenics sodium dodecy sulfate-polyacrylamide gel electrophoresis, prion protein identification, 32, 142, 150, 152 species barriers, to prion proteins in transmissible spongiform encephalopathies conformation analysis, 123–124 conversions, 156–159 disease host diversity, 13–17, 156–159 three-dimensional structure, 74–78 transgenic studies, 288–295 artificial prions, 289–290 bovine to human transmission, 294–295 bovine to murine transmission, 292–294 human to murine transmission, 290–292 mechanisms, 288 murine to hamster transmission, 288–289 spontaneous disease formation familial infection, 18–19 transgenic studies, 286–288 Ssa1, PSI metabolism regulation, 358–359 Ssb, PSI metabolism regulation, 359–360 SUP35 modeling in vitro, 346–347 prion evidence overexpression induction, 339 PSI link, 338–339 self-assembly, 346–347 sequence elements, 342–344 translation termination efficiency, 318 SUP35p, PSI relationship, 319

superoxide dismutase cellular stress resistance, 229–230, 300 copper binding, 10 neurodegeneration role, 300 Syrian hamster prion protein globular domain three-dimensional structure, 71, 74, 176 transgenic species barrier studies, 288–289

T therapeutic drugs, see drug research thermodynamics, recombinant prion protein folding stability, 96–101 toxicity, of prion proteins, 20, 146, 187–195, 300–302 transduction, receptor-mediated signal transduction, 233–235 transgenics prion protein structure influence, 273–304 disease propagation conditional Prnp knockout, 275–280 neurografted PrP-producing cells in null mice, 278–279 spontaneous disease, 286–288 gene ablation effects, 275–278 gene structure and function, 280–284 disease link, 283–284 internal deletions, 281–283 N-terminal deletions, 280–281 incubation studies, 295–297 molecular basis of prion strains, 297–302 description, 297 neurodegeneration mechanisms, 300–302 strain types, 297–300 overview, 273–275, 304 species barrier studies, 288–295 artificial prions, 289–290 bovine to human transmission, 294–295 bovine to murine transmission, 292–294 human to murine transmission, 290–292 mechanisms, 288 murine to hamster transmission, 288–289

404

SUBJECT INDEX

transgenics (continued) topology studies, 284–286 transgene vector considerations, 302–303 transmissible mink encephalopathy, description, 2 transmissible spongiform encephalopathies, see also specific diseases biosynthesis of prion proteins cellular biology cell culture models, 212–215 membrane attachment mechanisms, 216–218, 231–235 posttranslational cleavage, 222 production kinetics, 141, 218–220 subcellular formation site, 220–222 subcellular localization, 215–216 topological forms, 284–286 overview, 4–6, 203–204, 222 disease hosts species, 1–2 species barriers, see species barriers virus pathogenesis, 13 disease mechanisms, 2–3, 19–20, 286–288 disease strains, 10–13 conformation analysis, 124–125 prion protein conversions and interactions, 159–162 prion protein diversity, 10–13 familial infection mechanisms, 19–20 mutant PrP properties, 19, 125–128 mutant PrP-sen versus PrP-res, 19 spontaneous PrP-res formation, 18–19 glycosylation, 10, 12–13, 207–208 neurotoxicity of prion proteins, 20, 146, 187–195 overview, 1–3, 20–21, 55, 108 pathogenesis cell surface receptors role cadherins, 242–243 function, 231–235 37 kDa laminin receptor precursor, 230, 237–242 66 kDa membrane protein, 235–236 incubation period, 295–297 prion peptide fragment studies, 185–196 astroglial cultures, 194–196 cell membrane interactions, 185–187

human disease-associated mutations, 193–194 microglial cultures, 194–196 neuronal culture analysis, 187–194 (PrP 106-126), 187–193 yeast prions compared, 327–328 prion protein conversions and interactions anti-TSE drug research, 162–163 disease strains, 159–162 genetic link, 283–284, 338 infectivity, 162 mechanisms, 109, 129, 139–141 PrP deletion forms, 145 PrP-res folding and unfolding, 141–145 PrP-res formation, 145–148 PrP-sen-PrP-res interactions, 159–162 PrP structural diversity, 145–146 protein-only hypothesis, 83, 96, 248, 275 resistance, 9, 12–17 spontaneous disease formation, 18–19, 286–288 therapeutic drugs, 255–262 amphotericin B, 163, 257–260 anthracycline 4′-iodo-4′-deoxydoxorubicin, 257, 259, 261 carrageenan, 255 clusterin, 260–261 Congo red, 163, 257–259 Cp-60, 257, 261 Cp-62, 257, 261 dapsone, 260–261 dextran sulfate, 163, 255–256, 258 flurpirtin, 260–261 heparin, 230, 255 heteropolyanion (23), 163, 255–256 iPrP13, 257, 261 MS-8209, 257–260 overview, 230–231 pentosan polysulfate, 162, 255–256, 258–259 phtalocyans, 257, 261 polyanions, 255–258 polyene antibiotics, 258–260 porphyrins, 163, 257, 261 pythalocyanines, 163 three-dimensional structural change effects, 20, 56 trimethylamine-N-oxide, molecular chaperone role, 248

SUBJECT INDEX

two-state folding model, recombinant prion protein, 86-87,90-91

U URE2, nitrogen catabolite repression, 316–318 URE2p amyloid formation in vitro, 323–325 prion domains, 319–321 protease resistance, 322–323 URE3 relationship, 319, 322–323, 329 URE3 inheritance, 337 nitrogen catabolite repression, 316–318 prion evidence, 321–322, 329 URE2p relationship, 319, 322–323, 329

Y yeast prions, 313–329 genetic criteria, 314–316 gene mutation link to phenotypes, 315–316 overproduction link to prion emergence, 315 reversible curability, 314–315 het-s, 325–327, 329 nitrogen catabolite repression, 316–318 overview, 313–314, 329 Podospora anserina prions amyloidoses implications, 328–329 function, 325–327 PSI description, 335–336, 360 inheritance contributions from N region, 344–345 nonapeptide repeats, 344–345 N-terminus glutamines, 345 in vitro modeling, 346–352 alternate protein state replication, 347–350 N mutation biochemical effects, 350–352 SUP35 self-assembly, 346–347

405

metabolism regulation by molecular chaperones, 352–360 Hsp70, 354 Hsp104, 353–359 inheritance role, 356–357 maintenance role, 356–357 models, 357–358 Ssa1, 358–359 Ssb, 359–360 phenotype model, 341–342 prion evidence, 338–341 alternative phenotype associations, 339 alternative physical state replication, 339–341 SUP35 link, 338–339 SUP35 overexpression induction, 339 prion hypothesis formulation, 336–338 SUP35p relationship, 319 translation termination efficiency, 318 SUP35 modeling in vitro, 346–347 prion evidence overexpression induction, 339 PSI link, 338–339 self-assembly, 346–347 sequence elements, 342–344 translation termination efficiency, 318 SUP35p, PSI relationship, 319 translation termination efficiency, 318 TSE prions compared, 327–328 URE2, nitrogen catabolite repression, 316–318 URE3 inheritance, 337 nitrogen catabolite repression, 316–318 prion evidence, 321–322, 329 URE2p relationship, 319, 322–323, 329 URE2p amyloid formation in vitro, 323–325 prion domains, 319–321 protease resistance, 322–323 URE3 relationship, 319, 322–323, 329

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