Handbook of Neurochemistry and Molecular Neurobiology. Neural Protein Metabolism and Function

Handbook of Neurochemistry and Molecular Neurobiology Neural Protein Metabolism and Function Abel Lajtha (Ed.) Handb...

Author: Abel Lajtha | Naren Banik

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Handbook of Neurochemistry and Molecular Neurobiology Neural Protein Metabolism and Function

Abel Lajtha (Ed.)

Handbook of Neurochemistry and Molecular Neurobiology Neural Protein Metabolism and Function Abel Lajtha & Naren Banik

With 102 Figures and 73 Tables

Editor Abel Lajtha Director Center for Neurochemistry Nathan S. Kline Institute for Psychiatric Research 140 Old Orangeburg Road Orangeburg New York, 10962 USA Volume Editors Abel Lajtha Director Center for Neurochemistry Nathan S. Kline Institute for Psychiatric Research 140 Old Orangeburg Road Orangeburg New York, 10962 USA

Naren Banik Professor Department of Neurosciences Division of Neurology Medical University of South Carolina 96 Jonathan Lucas Street Suite 309 Charleston, SC 29425 USA

Library of Congress Control Number: 2006922553 ISBN: 978‐0‐387‐30346‐8 Additionally, the whole set will be available upon completion under ISBN: 978‐0‐387‐35443‐9 The electronic version of the whole set will be available under ISBN: 978‐0‐387‐30426‐7 The print and electronic bundle of the whole set will be available under ISBN: 978‐0‐387‐35478‐1 ß 2007 Springer ScienceþBusiness Media, LLC. All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC., 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. springer.com Printed on acid‐free paper

SPIN: 11416678 2109 ‐ 5 4 3 2 1 0

Table of Contents

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix

1

Regulation of Protein Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 M. Salinas . J. Burda

2

Cell Adhesion Molecules of the Immunoglobulin Superfamily in the Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 P. S. Walmod . M. V. Pedersen . V. Berezin . E. Bock

3

Neurosecretory Protein Trafficking and Dense‐Core Granule Biogenesis in Neuroendocrine Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 T. Kim . M. Gondre´‐Lewis . I. Arnaoutova . N. Cawley . Y. Peng Loh

4

Protein Synthesis at Synaptic Sites on Dendrites . . . . . . . . . . . . . . . . . . . . . . 169 O. Steward

5

Emerging Functions of the ‘‘Ca2+ Buffers’’ Parvalbumin, Calbindin D-28k and Calretinin in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 B. Schwaller

6

Actin, Actin‐binding Proteins and Myosins in Nervous System . . . . . . . . . . . 223 R. Ishikawa

7

Transglutaminase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 243 A. J. L. Cooper . S.‐Y. Kim

8

Protein Alterations in Mental Retardation . . . . . . . . . . . . . . . . . . . . . . . . . . . 259 M. A. Junaid . W. T. Brown

9

Protein Sulfation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283 S. Hemmerich

10

Protein Folding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303 A. Szila´gyi . J. Kardos . S. Osva´th . L. Barna . P. Za´vodszky

#

Springer-Verlag Berlin Heidelberg 2007

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Table of Contents

11

Neurodegenerative Dementias Involving Aberrant Protein Aggregation . . . 345 T. Kudo . M. Takeda

12

Polyglutamine Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 355 H. Okazawa

13

The Proteasome, Protein Aggregation, and Neurodegeneration . . . . . . . . . . 371 S. Wilk

14

Serine Proteases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 409 J. To´th . P. Medveczky . L. Szila´gyi . L. Gra´f

15

Calpain as a Target for Prevention of Neuronal Death in Injuries and Diseases of the Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 445 S. K. Ray . M. K. Guyton . E. A. Sribnick . N. L. Banik

16

Secretase Processing of Amyloid Precursor Protein (APP) and Neurodegeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 469 N. Marks . M. J. Berg

17

The Biology of Caspases in Central Nervous System Trauma . . . . . . . . . . . . . 515 M. L. McEwen . J. E. Springer

18

Microglial Proteases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 551 H. Nakanishi

19

MMPs and Other Matrix‐Degrading Metalloproteinases in Neurological Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 565 P. E. Gottschall . K. Conant

20

Protease Inhibitors and their Involvement in Neurological Disorders . . . . . . 591 B. Tizon . E. Levy

21

Neuropeptidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 625 K.‐S. Hui

22

Ubiquitination and Proteasomal Protein Degradation in Neurons . . . . . . . . . 653 L. Klimaschewski

23

Protease Activity in the Aging Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 663 D. A. Gray

24

From Concept to Potential Therapeutics: Neuroprotective Peptides . . . . . . . 673 I. Gozes . J. Tiong Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 689

Preface

This volume discusses aspects of neural protein metabolism. Although this subject is clearly very important and has numerous functional aspects, it did not always get the attention it clearly deserves. Perhaps there are many explanations for researchers hesitating in the past to focus on protein metabolism. Among the reasons, it must be that it was thought there is little or no regeneration in the brain; hence, its proteins must be stable. Stability, it was thought, is required in an organ that serves as a depository of information—the storage of memory. The structural and functional complexities and heterogeneity of brains may also have indicated that this organ compared to other tissues or even microbial systems may represent too much of a challenge. Technical difficulties also seemed to be discouraging. We have made significant progress, but still do not know much. For example, we now realize the multiple functions and activities of proteases in the nervous system; still, their activity, their specificity, and their controls in the living tissue remains a challenge to approach experimentally. The chapters in this volume clearly show that protein metabolism is very much ‘‘alive and active’’ in the nervous system and represents an essential part of brain functions. Proteins can and do undergo changes— modifications, partial or complete breakdown, structural and physical changes, and translocations—and are mostly in a dynamic state. It is true that the differences of processes between those in the living body and those in isolated systems—even complex isolated systems—are very large for proteins. Still, proteins are essential components and participate in practically all the processes in the nervous system, just as much as in other processes throughout living organisms. As proteins represent and are responsible for a great variety of functions and structures, their activity is complex. It is likely that the function and metabolism of a protein is different depending on its location. Clearly, some aspects of protein metabolism are not specific for the nervous system, but as they affect neural function, it is important to understand specifically their role in the nervous system. The many important aspects are clearly illustrated in this volume. It is as it should be focused on specific neural aspects and general aspects are left to other textbooks. Neuroscience has advanced during the past decades probably more than most biosciences; it represents a very exciting field. Probably no other field has the potential of influencing the future of mankind than studies of processes that underlie human behavior—all aspects: learning, memory, aggression, aging, and perhaps creativity. In some respects, it represents the ultimate challenge—scientifically as well as ethically. As our knowledge advances, our understanding of the functions of proteins represents an important segment. The fact that we begin to understand the role of proteins in neural and behavioral pathology represents an exciting opportunity for novel therapeutical approaches—some of which are discussed in this volume. The work described here should give us all hope of further advancing our knowledge and our possibility to cure mental and neurological illnesses in the not too distant future. This volume, as the others in the Handbook, is planned to be added to and brought up-to-date as new information is available and willing contributors can join. The field of neural proteins is large and full of possibilities, and we can expect rapid advance. Abel Lajtha and Naren Banik

Contributors

I. Arnaoutova Section on Cellular Neurobiology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD 20892, USA N. L. Banik Department of Neurosciences, Medical University of South Carolina, Charleston, SC 29425, USA L. Barna Institute of Enzymology, Hungarian Academy of Sciences, Budapest, Hungary

K. Conant Department of Neurology, Johns Hopkins University School of Medicine, Baltimore, Maryland, USA A. J. L. Cooper Departments of Biochemistry and Neurology and Neurosciences, Weill Medical College of Cornell University, 1300 York Avenue, New York, NY 10021, USA; and Burke Medical Research Institute, 785 Mamaroneck Avenue, White Plains, NY 10605, USA

V. Berezin Protein Laboratory, University of Copenhagen, Panum Institute, Blegdamsvej 3C, bld. 6.2., 2200 Copenhagen N, Denmark

M. Gondre´‐Lewis Section on Cellular Neurobiology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD 20892, USA

M. J. Berg Department of Psychiatry, New York University School of Medicine, Nathan S. Kline Institute for Psychiatric Research, Orangeburg, NY 10962, USA

P. E. Gottschall Department of Pharmacology and Therapeutics, University of South Florida College of Medicine, Tampa, Florida, USA

E. Bock Protein Laboratory, University of Copenhagen, Panum Institute, Blegdamsvej 3C, bld. 6.2., 2200 Copenhagen N, Denmark

I. Gozes Department of Clinical Biochemistry, Sackler Faculty of Medicine, Tel Aviv University, Tel Aviv 69978, Israel and Allon Therapeutics, Inc. Vancouver, BC V6J 4M6, Canada

W. T. Brown NYS Institute for Basic Research in Developmental Disabilities, 1050 Forest Hill Road, Staten Island, NY 10314, USA J. Burda Institute of Neurobiology, Slovak Academy of Sciences, Sˇolte´sovej 4, 040 01 Kosˇice, Slovakia N. Cawley Section on Cellular Neurobiology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD 20892, USA

#


L. Gra´f Protease Research Group of the Department of Biochemistry, Eo¨tvo¨s Lorańd University, Pa´zmańy Pe´ter st. 1/C, Budapest, H–1117, Hungary D. A. Gray Ottawa Health Research Institute, 501 Smyth Rd, Ottawa ON K1H 8L6, Canada M. K. Guyton Department of Neurosciences, Medical University of South Carolina, Charleston, SC 29425, USA

x

Contributors S. Hemmerich Y’s Therapeutics Inc., Burlingame, CA 94010, USA K.‐S. Hui Peptide Research Laboratory, Neurochemistry Division, Nathan S. Kline Research Institute for Psychiatric Research, Orangeburg, NY 10962, USA, and Department of Psychiatry, New York University School of Medicine, New York, NY, USA R. Ishikawa Department of Molecular and Cellular Pharmacology, Gunma University Graduate School of Medicine, Maebashi, Gunma 371–8511, Japan M. A. Junaid NYS Institute for Basic Research in Developmental Disabilities, 1050 Forest Hill Road, Staten Island, NY 10314, USA J. Kardos Department of Biochemistry, Eo¨tvo¨s Lorańd University, Budapest, Hungary T. Kim Section on Cellular Neurobiology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD 20892, USA S.‐Y. Kim Departments of Neurology and Neurosciences, Weill Medical College of Cornell University, 1300 York Avenue, New York, NY 10021, USA; and Burke Medical Research Institute, 785 Mamaroneck Avenue, White Plains, NY 10605, USA L. Klimaschewski Department of Anatomy and Histology, Division of Neuroanatomy Medical University of Innsbruck, Muellerstrasse 59, A–6020 Innsbruck, Austria

M. L. McEwen University of Kentucky Medical Center and the Cardinal Hill Rehabilitation Hospital, Departments of Physical Medicine and Rehabilitation, and Anatomy and Neurobiology, 800 Rose St., MN225, Lexington, KY 40536–0298, USA N. Marks Center for Neurochemistry, Department of Psychiatry, New York University School of Medicine, Nathan S. Kline Institute for Psychiatric Research, Orangeburg, NY 10962, USA P. Medveczky Protease Research Group of the Department of Biochemistry, Eo¨tvo¨s Lorańd University, Pa´zmańy Pe´ter st. 1/C, Budapest, H–1117, Hungary H. Nakanishi Laboratory of Oral Aging Science, Faculty of Dental Sciences, Kyushu University, Fukuoka 812–8582, Japan H. Okazawa Department of Neuropathology, Tokyo Medical and Dental University, Japan Science and Technology Agency (JST), PRESTO, Japan S. Osva´th Department of Biophysics and Radiation Biology, Semmelweis University, Budapest, Hungary M. V. Pedersen Protein Laboratory, University of Copenhagen, Panum Institute, Blegdamsvej 3C, bld. 6.2., 2200 Copenhagen N, Denmark Y. Peng Loh Section on Cellular Neurobiology, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD 20892, USA

T. Kudo Department of Psychiatry, Osaka University Graduate School of Medicine, D3, 2–2, Yamadaoka, Suita, Osaka 565–0871, Japan

S. K. Ray Department of Neurosciences, Medical University of South Carolina, Charleston, SC 29425, USA

E. Levy Departments of Psychiatry and Pharmacology, New York University School of Medicine, and Nathan Kline Institute, Orangeburg, New York 10962, USA

M. Salinas Servicio de Bioquı´mica‐Investigacioń, Hospital Ramoń y Cajal, Ctra. Colmenar Km 9, 28034 Madrid, Spain.

Contributors B. Schwaller Division of Histology, Department of Medicine, University of Fribourg, chemin du Museé 14, 1700 Fribourg, Switzerland J. E. Springer University of Kentucky Medical Center and the Cardinal Hill Rehabilitation Hospital, Departments of Physical Medicine and Rehabilitation, and Anatomy and Neurobiology, 800 Rose St., MN225, Lexington, KY 40536–0298, USA E. A. Sribnick Department of Neurosciences, Medical University of South Carolina, Charleston, SC 29425, USA O. Steward Reeve‐Irvine Research Center, and Departments of Anatomy & Neurobiology, Neurobiology & Behavior, and Neurosurgery College of Medicine, University of California at Irvine, Irvine, CA 92697, USA A. Szila´gyi Institute of Enzymology, Hungarian Academy of Sciences, Budapest, Hungary L. Szila´gyi Protease Research Group of the Department of Biochemistry, Eo¨tvo¨s Lorańd University, Pa´zmańy Pe´ter st. 1/C, Budapest, H–1117, Hungary M. Takeda Department of Psychiatry, Osaka University Graduate School of Medicine, D3, 2–2, Yamadaoka, Suita, Osaka 565–0871, Japan

J. Tiong Department of Clinical Biochemistry, Sackler Faculty of Medicine, Tel Aviv University, Tel Aviv 69978, Israel and Allon Therapeutics, Inc. Vancouver, BC V6J 4M6, Canada B. Tizon Departments of Psychiatry and Pharmacology, New York University School of Medicine, and Nathan Kline Institute, Orangeburg, New York 10962, USA J. To´th Protease Research Group of the Department of Biochemistry, Eo¨tvo¨s Lorańd University, Pa´zmańy Pe´ter st. 1/C, Budapest, H–1117, Hungary P. S. Walmod Protein Laboratory, University of Copenhagen, Panum Institute, Blegdamsvej 3C, bld. 6.2., 2200 Copenhagen N, Denmark S. Wilk Sherwin Wilk, Department of Pharmacology and Biological Chemistry, Mount Sinai School of Medicine, One Gustave L. Levy Place, New York, NY 10029, USA P. Za´vodszky Institute of Enzymology, Hungarian Academy of Sciences, Budapest, Hungary

xi

1

Regulation of Protein Metabolism

M. Salinas . J. Burda

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

2 2.1 2.1.1 2.1.2 2.2 2.2.1 2.2.2 2.2.3

Steps in Protein Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Initiation Step . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Initiation Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structural Elements of the mRNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elongation and Termination Steps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elongation Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Termination Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of Elongation and Termination Steps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3 3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.1.5 3.1.6 3.2 3.2.1 3.2.2 3.2.3 3.2.4 3.2.5

Mechanisms of General and Gene‐Specific Translational Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 General Translational Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 eIF2a Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 eIF2B Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 4E‐BP1 Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 eIF4E Levels and Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 eIF4G Cleavage and Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Translational Regulation by Other Initiation Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Gene‐Specific Translational Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Control at 50 ‐Oligopyrimidine Tract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Control at Internal Ribosome Entrance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Upstream Open Reading Frames in the 50 ‐UTR: Reinitiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Control by Cis‐Acting Elements Activated by Trans‐Acting Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 Control by MicroRNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15

4 4.1 4.2

Cerebral Ischemia‐Induced Downregulation of Translation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 Experimental Models of Cerebral Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15

5 5.1 5.2 5.2.1 5.2.2 5.3 5.3.1 5.3.2 5.4 5.4.1 5.4.2

Regulation of Protein Synthesis During Cerebral Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ischemic Period . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Early Reperfusion Period . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Initiation Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Endoplasmic Reticulum Dysfunction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Late Reperfusion Period . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Initiation Factors and Protein Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . eIF4G and Calpain Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Translation Control in Delayed Neuronal Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Control of Protein Expression: GluR2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Inhibition by Calpain Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

#


4 4 4 6 7 7 7 8

16 16 17 17 19 19 19 20 21 22 23

2

1

Regulation of protein metabolism

5.5 Translational Control in Ischemic Preconditioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 5.5.1 Improvement of Initiation at Early Reperfusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 5.5.2 Control of Survival Proteins: Expression of GADD34 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24 6

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24


1

Abstract: Translation is a sophisticated and complex mechanism requiring extensive biological machinery and subjected to an extremely fine regulation. Given that proteins account for a large fraction of biological macromolecules, a large proportion of the resource of cells is devoted to translation. Translation can be broken into three stages: initiation, elongation, termination. Regulation of translation can be exerted at many levels but in the first step, structural features in mRNAs in concert with the interactions among the initiation factors and other regulatory proteins, offer the possibility to regulate protein synthesis in rapid response to external stimuli, without invoking nuclear pathways for mRNA synthesis. The range of biological process that involve translational control of gene expression is expanding, in the brain, recent reports have provided evidences of the importance of translational control in process such us ischemia and reperfusion, neuronal plasticity and memory. List of Abbreviations: AMPA, alpha-amino-3-hydroxy-5methyl-4-isoxazolepropionic acid; ATF4, activating transcription factor 4; AUG, initiation codons; Cdk5, cyclin-dependent kinase 5; 4EBPs, initiation factor 4E binding protein family; eEFs, eukaryotic elongation factors; eIFs, eukaryotic initiation factors; eIF2α, the α subunit of initiation factor 2; eIF2(αP), the phosphorylated form of eIF2α; ER, endoplasmic reticulum; ERK, extracellular signal-regulated kinase; GADD34, growth arrest and DNA damage protein 34; GCN2, general control nonderepressible-2 kinase; GluR2, glutamate receptor 2 subunit of AMPA receptor; GRP78, glucose-regulated protein of 78 kDa; GSK-3, glycogen synthase kinase-3; HRI, heme-regulated inhibitor; IRES, internal ribosome entry sites; Met-tRNAi, initiator methionyl transfer RNA; m7G cap, 50 -m7G(50 )ppp(50 )N cap structure; Mnk, MAP kinase-interacting kinase; mTOR, target of rapamycin; NR2A, the N-metyl-D-aspartate (NMDA) receptor subunit 2A; uORF, upstream open reading frame; PABP, poly(A)-binding proteins; PKR, the double-stranded RNA-dependent kinase; PERK, mammalian ER resident kinase; PI3K/Akt-PKB, phosphatidylinositol-3-kinase/phosphatidylinositol-3-kinase-protein B; poly (A), 3´-polyadenylated tail; PP1c, catalytic subunit of protein phosphatase 1; S6K1/2, ribosomal protein S6 kinase 1/2; 5´TOP, terminal oligopyrimidine tracts; UPR, unfolded protein response; UTR, untranslated region

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Introduction

Given the importance that proteins occupy among the molecules crucial for life processes, a large proportion of resources within cells is devoted to protein synthesis. The translation of mRNAs into protein products occupies a position in the middle of a complex pathway that begins with transcription, continues with messenger RNA processing and transport, and ends with protein translocation, modification, folding, assembly, and degradation. Thus, translation constitutes a crucial point where gene expression can be regulated (Hershey, 1991; Mathews et al., 2000; Richter et al., 2001; Dever, 2002; Gebauer et al., 2004; Hay et al., 2004). Translational control is the mechanism used to regulate gene expression in many distinct biological situations such as early embryonic development, differentiation, and metabolism. In the brain, translational regulation plays an important role among the mechanisms that are implicated within the ischemic process. Since initiation factors are crucial components of protein synthesis machinery, those mainly involved in the ischemic process are reviewed in detail in this chapter. The presumption that the soma represented the primary site of protein synthesis in the neuron and that synapses depended on this synthesis for their function is no longer supported. Studies conducted in the last 10 years support the hypothesis that protein synthesis occurs in multiple subcellular compartments in the neuron, including dendrites, axons, and presynaptic nerve terminals (Kaplan et al., 2004). Many dendritically localized mRNAs encode for proteins that are critical for certain forms of synaptic plasticity (Paradies et al., 1997). Besides, the existence of functional protein synthesis machinery localized in the synaptodendritic compartment has been demonstrated (Job et al., 2001; Steward et al., 2001; Asaki et al., 2003). Compelling evidences now support that local protein synthesis within dendrites plays a key role in activity‐ dependent synaptic modifications (Huber et al., 2000; Steward et al., 2003; Kelleher et al., 2004a, b). Although we will focus on the control of translation in ischemia and reperfusion, we will briefly mention

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those particular factors and mRNA elements implicated in global and gene‐specific translation regulation in neuronal plasticity. Given the extraordinary complexity of the translational machinery and the multiple aspects of the ischemic process, references to the most recent published reviews covering each specific topic are made at each section. To supplement this brief survey and for a more comprehensive citation of primary research literature of ischemic process we also refer the reader to other chapters within this series handbook.

2

Steps in Protein Synthesis

Protein synthesis is a sophisticated multistep process requiring extensive biological machinery and a substantial input of energy. Ribosomes, large ribonucleoprotein assemblies acting in concert with a number of accessory factors, translate the genetic information contained in messenger RNA (mRNA) molecules. The dynamic process of mRNA translation is conveniently divided into three phases or steps: initiation, elongation, and termination (Hershey, 1991; Merrick et al., 1996). The initiation phase represents all processes required for the assembly of a ribosome with the unique initiator methionyl transfer RNA (Met‐tRNAi) in its peptidyl (P) site and locating the proper AUG (although other codons may be occasionally used) start codon on the mRNA. The polypeptide synthesis takes place during the elongation phase and peptide bond formation occurs on catalytic centers that are fundamentally formed by the ribosomal RNA (rRNA) of the large ribosomal subunit. When the ribosomes reach the stop codon, this signals termination comprising the release of the complete polypeptide and, presumably, the ribosome from the mRNA. The ribosome itself cannot perform these functions, which are facilitated by specific proteins referred as translation factors. This is particularly true during the initiation step in eukaryotes where both ends of mRNA, the 50 ‐m7G(50 )ppp(50 )N cap structure (m7G cap) and the 30 ‐polyadenylated (poly(A)) tail, as well as at least twelve initiation factors (eIFs) are required (Pain, 1996; Hershey et al., 2000; Preiss et al., 2003).

2.1 Initiation Step 2.1.1 Initiation Factors Formation of the 43S Complex In eukaryotes, binding of Met‐tRNAi to the 40S ribosomal subunit is primarily the responsibility of initiation factor 2, eIF2, which forms a stable ternary complex with GTP and Met‐tRNAi (> Figure 1-1). The ternary complex is delivered directly to the P site of the ribosome whereas the other aminoacyl‐tRNAs are delivered to the A site in elongating ribosomes. Four more factors help in this step, eIF1A and the multisubunit factor eIF3 generate a pool of free 40S ribosomal subunits, and the binding of the ternary complex to this ribosomal subunit is also aided by eIF1 and eIF5 (Hershey, 1991; Hinnebusch, 2000; Preiss et al., 2003). Formation of the 48S Complex Binding of the 40S·Met‐tRNAi·GTP·eIF2 (43S preinitiation complex) to the mRNA is promoted by the eIF4 factors and eIF3 (> Figure 1-1). The heterotrimeric complex eIF4F is composed of eIF4E, eIF4G, and the dead‐box RNA helicase eIF4A. eIF4E directly binds the m7G cap at the 50 ‐end of eukaryotic mRNAs. eIF4G is a multivalent adapter molecule whose N‐terminal harbors binding sites for eIF4E and the poly(A)‐binding proteins (PABP) that serve to latch it onto both RNA ends, implying the potential to pseudocircularize the mRNA (Hershey, 1991; Gingras et al., 1999; Preiss et al., 2003). The central third region of eIF4G interacts with eIF4A, mRNA, and eIF3, where eIF4A in conjunction with two binding proteins, eIF4B and eIF4H (an eIF4B homolog), is thought to unwind RNA secondary structures near the 5’‐end. This process is dependent on ATP hydrolysis but it is not known whether eIF4A or another ATPase facilitates the scanning process by melting mRNA secondary structures or by actively propelling the ribosome (Kozak, 1989; Gingras et al., 1999; Kozak, 1999). In mammals, at the extreme C‐terminal of eIF4G there is a region where eIF4E kinase, MAP kinase interacting kinase (Mnk1), binds.


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. Figure 1-1 Schematic representation of the translation initiation pathway. In a round of initiation, the successive assembly of the small (40S) ribosomal subunit, the ternary complex formed by eIF2, Met‐tRNAi and GTP, the mRNA, and the large (60S) ribosomal subunit, with the participation of distinct initiation factors, forms the 80S initiation complex on the mRNA required for protein synthesis. The ternary complex aided by eIF3, eIF1, eIF1A, and eIF5, binds to the small (40S) ribosomal subunit to yield the 43S preinitiation complex. This complex is recruited to the mRNA via the interaction of eIF3 with eIF4G in the eIF4F complex. The eIF4F contains eIF4E that directly binds to the 50 ‐cap and eIF4A, a helicase that unwinds the secondary structure of the mRNA during the scanning step. In the N‐terminal third, eIF4G binds the poly(A) tail of mRNA through interaction with PABP proteins and in the C‐terminal it harbors a binding site for Mnk1, the eIF4E kinase. The 43S preinitiation complex assisted by eIF1 and eIF1A then scans the mRNA in a 50 –30 direction until recognition of the first AUG codon, which involves base‐pairing with the anticodon in the loop of the Met‐tRNAi, yielding a stable 48S ribosomal complex. After the hydrolysis of GTP bound to eIF2, catalyzed by eIF5, several factors are released. A second GTPase, termed eIF5B, is required for the joining of the large subunit (60S), resulting in the formation of an 80S complex competent to catalyze the first peptide bond. ATP hydrolysis on eIF4A (stimulated by eIF4B) and the two GTP hydrolysis on eIF2 and eIF5B are shown in the outside of the complex for simplicity. The interactions among eIF1, eIF1A, eIF2, eIF5, and eIF3 in the 48S complex are only approximated and the process for 80S formation is not given in detail in this scheme. The recently identified eIF4H factor that stimulates the helicase activity of eIF4A is not included. From Cristina Martıń de la Vega (Ph.D. thesis: January 18, 2002 to be awarded)

Scanning and Assembly of the 80S Ribosome Once the 43S complex is assembled to the 50 ‐end of

the mRNA, forming the 48S complex, it initiates a linear movement, ‘‘scanning’’, along the mRNA 50 ‐untranslated region (50 ‐UTR) to recognize the appropriate start codon that in most mRNAs is the AUG triplet closest to the 50 ‐end. The proper recognition of this start codon depends on base pairing contacts with the anticodon of the Met‐tRNAi as well as factors eIF1, eIF1A, eIF2, and eIF5. At least in

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mammals, recognition of an AUG as a translation start codon also critically depends on its surrounding sequence (Kozak, 1991; Meijer et al., 2002; Preiss et al., 2003). In summary, the assembly of the 43S preinitiation complex at the 50 ‐cap structure of the mRNA is principally directed by the cap‐binding protein eIF4E and coordinated by eIF3 and eIF4G, which provide multiple contact points to other initiation factors, the ribosomal subunit, and the mRNA. Upon AUG codon recognition the codon–anticodon interaction probably triggers a conformational change in 40S‐subunit‐bound eIF2, which leads to GTP hydrolysis aided by the GTPase activating protein, eIF5, which is bound to the b‐subunit of eIF2. This is followed by the release of many or all the factors from the 40S, leaving the Met‐tRNAi in the P‐site base‐paired to the AUG start codon in the mRNA. It is catalyzed by eIF5B whose GTPase activity is activated by 60S subunit and more strongly by the combination of 40S and 60S subunits. GTP hydrolysis is required for eIF5B release and occurs after 80S complex formation (Pestova et al., 2000; Lee et al., 2002). This reaction completes the initiation phase and primes the ribosome to accept the first elongation aminoacyl‐tRNA. The pathway depicted in > Figure 1-1, where the 48S complex scans down the mRNA in a 50 –30 direction searching for the AUG start codon, applies to the majority of translation initiation events in eukaryotic cells. However, a few alternative mechanisms such as ribosome shunting and internal initiation have been reported and are discussed later (Carter et al., 2000; Jackson, 2000; Hellen et al., 2001; Preiss et al., 2003).

2.1.2 Structural Elements of the mRNA The different proteins that can be synthesized from one mRNA are marked by the choice of the translation initiation codon. This choice as well as the intrinsic translational efficiency is controlled by diverse structural features and regulatory sequences within the mRNA. These cis‐acting elements can be divided into two categories: those that act alone or with general translation factors; and those whose actions are mediated by additional specific trans‐acting factors. Cis‐Acting Elements An mRNA consists, in the 50 –30 direction, of the m G cap, the 50 ‐UTR, i.e., the 7

sequence between the cap and the initiation codon of the main open‐reading frame (ORF), the 30 ‐UTR, and a poly(A) tail. All these cis‐acting elements contribute significantly to translation efficiency, although the ORF itself is in general not involved (Kochetov et al., 1998; Geballe et al., 2000; Dever, 2002; Meijer et al., 2002; Gebauer et al., 2004) (> Figure 1-2). In fact, the m7G cap and the 30 ‐poly(A) tail found on most eukaryotic mRNAs synergistically enhance the translational efficiency of the mRNA (Preiss et al., 2003). In most cases, 50 ‐UTRs that enable efficient translation are short, have a low CG content and an optimal context of nucleotides flanking the initiation codon, are relatively unstructured, and do not contain upstream initiation codons (uAUGs). Interestingly, it has been demonstrated that these naturally ‘‘strong’’ mRNAs encode highly abundant cellular proteins such as actins involved in cytoskeleton . Figure 1-2 Schematic representation of structural elements of mRNA. An mRNA consists, in the 50 –30 direction, of the m7G cap, the 50 ‐untranslated region (50 ‐UTR), the main open reading frame (ORF), the 30 ‐UTR, and a poly(A) tail. All these cis‐elements contribute to translation efficiency, in fact, the presence of m7G cap, poly(A) tail, as well as a relatively unstructured 50 ‐UTR synergistically contribute to enhance translation. Secondary structures such as hairpins contribute to repress or block translation. The presence of internal ribosome entry sites (IRES), which contributes to cap‐independent translation, and terminal oligopyrimidine tracts (50 ‐TOP) or upstream open reading frames (uORFs), allow the translational control of diverse gene groups by distinct mechanisms. Cis‐ acting elements requiring trans‐acting factors usually regulate inhibitory mechanisms, but infrequently may activate translation


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formation, ribosomal proteins, translational machinery factors, and heat‐shock proteins (Kochetov et al., 1998; Schneider, 2000). On the contrary, 50 ‐UTRs with extensive secondary structure are translated less efficiently, because in order to allow ribosome access to the 50 ‐UTR these secondary structures need to be melted, possibly by eIF4F and eIF4B action. When eIF4F levels are low, the translation of these ‘‘weak mRNAs’’ encoding protein products that function in controlling cell growth and proliferation is impaired (Kozak, 1994; Gingras et al., 2004). All the other different elements present in the 50 ‐UTRs, including secondary or tertiary mRNA structures such as terminal oligopyrimidine (50 ‐TOP) tracts (see > Sect. 3.2.1) (Meyuhas, 2000), hairpins or pseudoknots, internal ribosome entry sites (IRES) (see > Sect. 3.2.2), uAUGs, and uORFs (see > Sect. 3.2.3), are very probably used to enable the translational control of several gene groups by distinct mechanisms. Cis‐Acting Elements Dependent on Trans‐Acting Factors Cis‐acting elements requiring trans‐acting factors

are also present throughout the mRNA and in general most of the regulatory mechanisms discovered so far are inhibitory. Ferritin was one of the first proteins recognized as being translationally regulated and mammalian cells use this mechanism to ensure that ferritin is expressed primarily when required for the sequestration of excess iron (Rouault et al., 2000). The iron‐responsive element (IRE) found in the 50 ‐UTR of ferritin mRNA is repressed by the trans‐acting iron repressor proteins IRP1 and IRP2. When cytosolic iron levels increase, iron binds to IRP1/2 and the proteins are released from the transcript (Hentze et al., 1996; Gebauer et al., 2004). The 30 ‐UTR is also a rich repository of cis‐acting elements modulated by trans‐acting factors, which determine among others, mRNA stability by the interaction between poly(A)‐binding protein (PABP) and the N‐terminal part of eIF4G (Sachs, 2000). Mammalian cells have evolved additional PABP‐interacting proteins, termed Paip1 and Paip2, that seem to act as translational activators or repressors respectively (Preiss et al., 2003). In addition, removal of the 30 ‐poly(A) tail initiates the degradation of mRNAs (Coller et al., 2004). Another cis‐acting element in the 30 ‐UTR, the cytoplasmic polyadenylation element (CPE), recruiting the trans‐acting factor, cytoplasmic‐polyadenylation element‐binding protein (CPEB), mediates processes such as mRNA transport, including targeting of mRNAs to dendrites in response to synaptic stimulation (Rook et al., 2000; Mendez et al., 2002; Huang et al., 2003) and translational activation (Groisman et al., 2000; Mendez et al., 2001).

2.2 Elongation and Termination Steps 2.2.1 Elongation Factors The elongation cycle of protein biosynthesis on the ribosome adds one amino acid at a time to a growing polypeptide according to the sequence of codons found in the mRNA. In general, the elongation step in eukaryotes is thought to be similar to, if not identical to, prokaryotes and it requires ancillary elongation factors (Andersen et al., 2003). The recruitment of the aminoacyl‐tRNA to the A site is catalyzed by eukaryotic elongation factor 1A (eEF1A) that forms the ternary complex aa‐tRNA·eEF1A·GTP. The peptidyl transferase center of the ribosome catalyzes the formation of a peptide bond between the incoming amino acid and the peptide found in the peptidyl‐tRNA binding site, P site. The subsequent translocation event is catalyzed by elongation factor 2 (eEF2). Both factors eEF1A and eEF2 are GTP‐binding proteins; the GTP is hydrolyzed during the events that they regulate and end up in an inactive form bound to GDP. The recycling of GDP by GTP is spontaneously done for eEF2, but eEF1A needs the help of another factor, the nucleotide exchange factor eEF1B (Merrick et al., 2000; Proud, 2000).

2.2.2 Termination Factors The presence of one of the three termination codons, UAA, UAG, or UGA in the A site of the ribosome signals the polypeptide chain release factors to bind and recognize the termination signal. In eukaryotes,

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translation termination is catalyzed by eRF1, which recognizes all three termination codons. When a stop codon is located in the ribosomal A site, polypeptide chain release factors are recruited to the ribosome to catalyze the hydrolysis of the ester bond between the final amino acid of the polypeptide chain and the tRNA, resulting in the release of a mature polypeptide chain. A second release factor, eRF3, stimulates eRF1 activity in a GTP‐dependent fashion (Hoshino et al., 1999; Welch et al., 2000).

2.2.3 Regulation of Elongation and Termination Steps Compelling evidences support that the elongation and termination steps of protein synthesis are also susceptible to regulation and several recent review articles cover the topic more comprehensively (Proud, 2000; Welch et al., 2000; Traugh, 2001; Proud, 2004). Cellular levels of elongation factors eEF1A and eEF2 are subject to translational control by the effect of certain hormones and changes in amino acid supply through the 50 ‐TOP sequences present in their mRNAs (Proud, 2000). The three elongation factors are phosphoproteins, and among other less characterized regulatory mechanisms phosphorylation of translation elongation factor, eEF2, seems to play a significant role in elongation step regulation. Concomitant to protein synthesis inhibition, eEF2 phosphorylation has been reported in neuronal cells exposed to stimuli known to raise intracellular Ca2þ levels (Marin et al., 1997; Alirezaei et al., 2001; Nairn et al., 2001) and in mammalian tissues subjected to mild ATP depletion (Browne et al., 2002; McLeod et al., 2002). Conversely, eEF2 dephosphorylation by protein phosphatase 2A (PP2A) induces protein synthesis reactivation (Browne et al., 2002; Horman et al., 2002; Yan et al., 2003). Translation termination efficiency is influenced by a number of trans‐acting factors and is now just beginning to be elucidated that it can act as a control point to regulate gene expression. Besides, it has been suggested that termination factors associated with the 50 ‐and 30 ‐ends of the mRNA interact to regulate both stability and translation efficiency of the transcript (Hoshino et al., 1999; Welch et al., 2000). The lack of correlation between eEF2 phosphorylation status and ischemia‐induced translation inhibition questions the participation of the elongation step within this process (Althausen et al., 2001; Garcıá et al., 2004a). Likewise, data concerning the potential relevance of termination step regulation during the ischemic process are not available. Due to space limitations, this chapter focuses on current knowledge regarding the initiation phase of translation, which is a major target for both global and mRNA‐specific translational regulation.

3

Mechanisms of General and Gene‐Specific Translational Control

Translational control is defined as a change in the rate (efficiency) of translation of one or more mRNAs, i.e., the number of complete protein products changes per mRNA molecule per unit time. It is generally believed that during protein synthesis the number of protein chains initiated is about the same as the number of proteins completed; consequently, under steady‐state conditions, the rate of initiation determines the rate of protein synthesis (Mathews et al., 2000). Structural features in mRNAs in concert with the interactions among the initiation factors facilitate the speed and accuracy of translation initiation and provide an opportunity for regulation.

3.1 General Translational Control Global regulation mainly occurs by the modification of translation initiation factors; many of them are phosphoproteins whose activities are regulated by specific kinases (Hershey, 1991; Proud, 1992; Pain, 1996; Roads, 1999). In general, growth stimulatory signals including hormones, growth factors, and mitogens lead to increased eIF4F complex formation and enhanced translation (Gingras et al., 1999; Roads, 1999; Lawrence Jr et al., 2001; Morley, 2001; Herbert et al., 2002; Hay et al., 2004). On the contrary, cellular


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stress conditions including iron deficiency, nutrient deprivation, heat shock, virus infection, changes in intracellular calcium, endoplasmic reticulum stress, and ischemia, inhibit global protein synthesis. Remarkably, most of these stress situations lead to the phosphorylation of the a‐subunit of eIF2 (Ochoa, 1983; Burda et al., 1994; Brostrom et al., 1998; Ron et al., 2000; Clemens, 2001; Deng et al., 2002).

3.1.1 eIF2a Phosphorylation The global control of initiation in most of the situations is commonly exerted on two sites, the first being the ternary complex formation between eIF2, GTP, and Met‐tRNAi. eIF2B is also required at this stage for the guanine nucleotide exchange on eIF2 to regenerate active eIF2·GTP (> Figure 1-1). eIF2 consists of three different subunits, a, b and g and phosphorylation of the a‐subunit, eIF2(aP), blocks global mRNA translation (Ochoa, 1983; Cale´s et al., 1985; Colthurst et al., 1987). Because the cellular ratio of eIF2B/eIF2 is about 1:3 to 5 (approximately 1:5 for the brain, (Alca´zar et al., 1995)), 20–30% of phosphorylated eIF2a acting as a competitive inhibitor of eIF2B activity may completely block initiation in most cells (Ochoa, 1983; Pain, 1996). eIF2a is phosphorylated at residue Ser51 by a family of very specific kinases, which includes the heme‐regulated inhibitor (HRI), expressed predominantly in erythroid cells, the double‐ stranded RNA‐dependent kinase (PKR), the general control nonderepressible‐2 kinase (GCN2, mGCN2), and the recently characterized mammalian ER resident kinase (PERK) (de Haro et al., 1996; Shi et al., 1998; Berlanga et al., 1999; Harding et al., 1999). The four kinases share extensive homology in the kinase catalytic domains, but the regulatory domains and, consequently, the regulatory mechanisms of each of them are very different. Hence, the reversible phosphorylation of eIF2a is a modification used for translational control in response to different environmental stresses that activate the corresponding kinase; i.e., iron deficiency, heat shock, or oxidative stress activates HRI (Chen, 2000); interferon, antiproliferative responses, and cell death program modulate PKR activity (Kaufman, 2000); and amino acid deprivation, UV irradiation, or ER stress activate mGCN2 (Deng et al., 2002) or PERK (Ron et al., 2000; Harding et al., 2002), respectively. Reduction of eIF2·GTP levels increases the time required for the scanning ribosomes to become competent enough to reinitiate translation and, hence, it limits initiation events on all cellular mRNAs very efficiently and rapidly and saves energy that can be diverted to other vital cellular processes.

3.1.2 eIF2B Phosphorylation eIF2B factor is a heteropentameric protein composed of subunits a to e with masses ranging from 26 to 82 kDa (Alca´zar et al., 1995; Proud, 2001). eIF2B activity can be mainly regulated by two mechanisms. First, phosphorylation of eIF2a inhibits eIF2B because eIF2(aP) is a competitive inhibitor of eIF2B in the GDP/ GTP exchange reaction (see > Sect. 3.1.1). Secondly, eIF2B activity can be regulated, in mammalian cells, in response to a variety of stimuli that share the common feature of activating translation (Kimball et al., 1996; Munõz et al., 1998; Quevedo et al., 2000; Wang et al., 2001). Four kinases have been described to phosphorylate the e subunit of eIF2B (eIF2Be); they are casein kinases (CK) 1 and 2, glycogen synthase kinase‐3 (GSK3), and dual specificity tyrosine‐phosphorylated and ‐regulated kinase (DYRK). Phosphorylation by CK2 enhances eIF2B activity, whereas phosphorylation by GSK3 has an inhibitory effect and requires previous eIF2Be phosphorylation by DYRK (Wang et al., 2001a; Woods et al., 2001). Upon treatment with different growth factors, activation of eIF2Be correlates with the activation of the phosphatidylinositol‐3‐kinase/phosphatidylinositol‐3‐kinase‐protein B (PI3K/Akt‐PKB) pathway, inactivation of GSK3, dephosphorylation of the GSK3 site on eIF2Be and increased global translation (Quevedo et al., 2000, 2003; Proud, 2001). In our laboratory, we have demonstrated that insulin‐like growth factor 1 also induces eIF2B activation in neurons via a new pathway mediated by protein phosphatase1/mitogen‐ activated protein kinase (PP1/MAPK) signaling, and our data suggest that, at least in neuronal cells, PP1 may be a physiological eIF2B phosphatase (Quevedo et al., 2003).

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3.1.3 4E‐BP1 Phosphorylation The second initiation site subjected to fine control is the recruitment of mRNA to a ribosome, a complex process which facilities its transfer to the 43S initiation complex, resulting in the 48S initiation complex. As shown in > Figure 1-1, this step requires the participation of eIF4‐family‐like initiation factors, eIF4F and eIF4B. The integrity of the eIF4F cap‐binding complex is regulated by eIF4E availability that is controlled by 4E binding protein family (4E‐BPs), a family of translational repressor proteins (Pause et al., 1994; Gingras et al., 1999). Nonphosphorylated 4E‐BPs compete with eIF4G for a common site on eIF4E, forming a complex that acts as a competitive inhibitor of eIF4G binding to eIF4E and prevents eIF4F formation. 4E‐BPs are phosphorylated at least on seven residues Thr37, Thr46, Ser65, Thr70, Ser83, Ser110, and Ser112 numbered according to the human sequence. The first five phosphorylation sites are conserved phylogenetically among all species but the last two only exist in 4E‐BP1. This isoform has been used in most of the studies, but it is not the most abundant in all tissues; in the brain, 4E‐BP2 is the most abundant isoform. It is postulated that 4E‐BP1 phosphorylation proceeds in an ordered and hierarchical manner but there is no general agreement on the role of the different phosphorylation events in the release of 4E‐BP1 from eIF4E (Beretta et al., 1996; Gingras et al., 1999; Mothe‐Satney et al., 2000; Gingras et al., 2001; Herbert et al., 2002; Hay at al., 2004). Many different extracellular stimuli including hormones, growth factors, and mitogens eliciting an increase in translation induce 4E‐BP hyperphosphorylation; conversely, nutrient deprivation and certain stress conditions decrease 4E‐BP1 phosphorylated levels. Several kinases are capable of phosphorylating 4E‐BP1 in vitro and rapamycin blocks both 4E‐BP1 phosphorylation and cap‐dependent translation in vivo, indicating that 4E‐BP1 is a downstream effector of mTOR (target of rapamycin). mTOR is a kinase that has emerged as a major effector of cell growth and proliferation via the regulation of protein synthesis and immunoprecipitates of mTOR phosphorylate the two priming sites in mammalian 4E‐BP1, Thr37 and Thr46, in vitro (Schmelzle et al., 2000; Gingras et al., 2004). However, it is not clear yet whether the intrinsic kinase activity of mTOR is sufficient for its full activity in vivo. It may act as a scaffold for other proteins with catalytic activity including other kinases and also phosphatases that may be responsible for its overall activity in vivo (Schmelzle et al., 2000; Lawrence Jr et al., 2001; Gingras et al., 2004; Hay et al., 2004). In hippocampal neurons, multiple forms of neuronal activity, including brain nerve growth factor treatment, excitatory synaptic activity, and membrane depolarization, stimulate translational efficiency in association with increased phosphorylation of eIF4E and 4E‐BP1/2 in an extracellular signal‐regulated kinase (ERK)‐dependent manner (Tang et al., 2002; Banko et al., 2004; Kelleher et al., 2004a,b). Both phosphorylations are inhibited in the hippocampus of transgenic mice expressing a dominant‐negative ERK kinase, which exhibits selective defects in the translational component of L‐LTP and memory consolidation (Kelleher et al., 2004a). Thus, 4E‐BP1 represents an excellent target for neuronal activity‐ dependent regulation of global mRNA translation in long‐lasting synaptic plasticity.

3.1.4 eIF4E Levels and Phosphorylation As a decreased availability of eIF4E downregulates translation, increased expression of eIF4E would be expected to increase eIF4F formation and increase translation. In fact overexpression of eIF4E in rodent cell lines causes malignant transformation and eIF4E is elevated in several human carcinomas (Clemens, 1999; De Benedetti et al., 1999; Martıń et al., 2000a; Wang et al., 2001b; Avdulov et al., 2004; Lynch et al., 2004; Rosenwald, 2004). It is now becoming evident that the main effect of increased eIF4E availability is the specific stimulation of those inefficiently translated mRNAs that contain excessive secondary structure in their 50 ‐UTR regions, requiring greater eIF4F complex levels to be translated. These ‘‘weak’’ mRNAs encode for regulatory proteins whose expression is low and are under stringent control, such as oncogenes and growth‐promoting proteins (Kozak, 1994; Kochetov et al., 1998; Gingras et al., 2004). A serum‐stimulated, rapamycin‐sensitive phosphorylation site in eIF4E has been found, Ser209, but the phosphorylation is not strictly required for eIF4E function and the precise mechanistic consequences of eIF4E phosphorylation are unclear (Gingras et al., 1999; Raught et al., 1999; Morley, 2001). The two


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different pathways that induce eIF4E phosphorylation, mitogen‐stimulated pathway acting through the ERKs and stress‐activated pathway through p38 mitogen‐activated protein kinase, p38 MAPK, converge at the common eIF4E kinase, Mnk1/2 (Fukunaga et al., 1997; Mahalingam et al., 2001). Very recently we have identified and characterized a splice variant of human Mnk1 gene (Mnk1b), which phosphorylates eIF4E in vitro and in vivo in a mitogen‐activated protein kinase‐independent manner (O’Loghlen et al., 2004). Mnk1 phosphorylates eIF4E both in vitro and in vivo but does not interact directly with the factor. Rather, Mnk1 binds to eIF4Gs and the interaction between eIF4E and eIF4G is required for eIF4E phosphorylation in vivo (Gingras et al., 1999; Pyronnet et al., 1999 Morley, 2001). Although phosphorylation of eIF4E generally correlates with increased translation rates (Gingras et al., 1999; Banko et al., 2004; Hay et al., 2004), recent studies have challenged this view (Knauf et al., 2001; McKendrick et al., 2001; Morley et al., 2002; Quevedo et al., 2002; Ueda et al., 2004).

3.1.5 eIF4G Cleavage and Phosphorylation Two isoforms of eIF4G, eIF4GI and eIF4GII, have been characterized, eIF4GI being the prototype form of the family; they exhibit moderate sequence conservation (46% identity in humans) but share similar overall biochemical activities (Gingras et al., 1999). Infection of cells with members of the picornavirus family results in a rapid shutoff of host cell protein synthesis, whereas poliovirus RNA translation is not affected. The poliovirus‐encoded protease 2A was found to be responsible for the specific cleavage of both eIF4GI and eIF4GII into two fragments that liberates the eIF4E‐binding site (N‐terminal fragment) from other functional regions, promoting the shutoff of cap‐dependent host protein synthesis. However, the larger C‐terminal fragment retains the ability to interact with eIF4A and eIF3 allowing IRES‐dependent translation of viral mRNAs (Gingras et al., 1999; Belsham et al., 2000; Pre´voˆt et al., 2003; Hay et al., 2004). The cleavage of eIF4GI and eIF4GII is a relatively early event in apoptosis that is mediated by caspase‐3 and yields discrete breakdown products termed FAGs (fragments of apoptotic cleavage of eIF4G), which retain binding sites for eIF4E, eIF4A, and eIF3 (Bushell et al., 1999, 2000; Morley, 2001; Pre´voˆt et al., 2003). Because the induction of apoptosis has been shown to be associated with a rapid, although incomplete, inhibition of global protein synthesis in several cell types, it is believed that FAGs may mediate cap‐ dependent as well as cap‐independent translation of specific mRNAs intimately associated with cell death (Pyronnet, 2000; Morley, 2001). This is in agreement with the fact that under severe apoptotic conditions, some cellular IRES‐driven mRNAs such as c‐myc, Apaf‐1, XIAP, and DAP‐5 continue to be translated (Pre´voˆt et al., 2003). Two different sets of phosphorylation sites have been identified in eIF4GI. The first contains three serum‐stimulated phosphorylation sites residing between the middle and the carboxyl‐terminal eIF4A‐ binding domains (Ser 1108, 1148, and 1192) and are modulated by the activity of the PI‐3K/Akt/mTOR kinase pathways. The second set, not wellcharacterized, is in the amino terminus end (Gingras et al., 1999; Raught et al., 2000). eIF4GII is phosphorylated to a lower extent than eIF4GI in a serum‐ or mitogen‐ independent way and calcium calmodulin kinase I (CaMKI) seems to be the kinase responsible for this phosphorylation both in vitro and in vivo (Qin et al., 2003). Nevertheless, the functional consequences of these phosphorylation events on eIF4G activity and how they may affect the translation initiation rate are unknown at this time.

3.1.6 Translational Regulation by Other Initiation Factors The potential role of other initiation factors such as eIF1, eIF1A, eIF3, eIF4A, eIF4B, eIF5, and eIF5B in translational control needs to be fully addressed. eIF3 is the largest of the mammalian initiation factors and consists of a complex of 11 subunits ranging from 35 to 170 kDa (Browing et al., 2001). The factor operates through its distinct subunits at different levels of the initiation pathway, including prevention of the association of 40S and 60S subunits, stabilization of ternary complex formation, and interaction with eIF4F‐mRNA to bring the mRNA to the 43S ribosomal complex (Hui et al., 2003; Preiss et al., 2003; Fraser et al., 2004;

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Nielsen et al., 2004). In addition, eIF3 seems to be required for encephalomyocarditis and hepatitis C virus IRES‐mediated translation (Theurl et al., 2004). Initiation factor eIF5 exists as phosphoprotein and the kinase responsible for its phosphorylation both in vitro and in vivo is casein kinase 2. Although binding of eIF2b to eIF5 seems to be essential for eIF5‐ promoted GTPase activity, phosphorylation of eIF5 does not seem to modify this activity in vitro (Majumdar et al., 2002). Treatment of cells with serum, insulin, or phorbol esters results in hyperphosphorylation of eIF4B and many kinases including ribosomal protein S6 kinase‐1 (S6K1), protein kinase C and A, casein kinase 1 and 2 are capable of phosphorylating eIF4B in vitro. It has been suggested that increased phosphorylation of eIF4B enhances eIF4B activity and the translation of mRNAs containing some degree of secondary structure (Dmitriev et al., 2003). It is becoming evident at this time that eIF4B is a physiologically relevant target of S6K1 in vivo (Raught et al., 2004). Very recent results suggest that the DEAD‐box RNA helicase Vasa (Vas), which is required for germ cell development and function, could work as a translational regulator of specific mRNAs through interaction with eIF5B (Johnstone et al., 2004).

3.2 Gene‐Specific Translational Regulation The mechanisms of global regulation by changes in the general translational machinery, as already discussed, may paradoxically have mRNA‐specific effects because, in general, weaker mRNAs seem to be more sensitive to perturbations in translation initiation efficiency and are downregulated when translation is impaired. In addition, another mechanism of gene‐specific translation regulation also involves the formation of regulatory protein complexes that recognize the particular cis‐elements present in the 50 ‐UTRs and 30 ‐UTRs of the target mRNA (Meyuhas, 2000; Hellen et al., 2001; Dever, 2002; Meijer et al., 2002; Gebauer et al., 2004).

3.2.1 Control at 50 ‐Oligopyrimidine Tract Many components of the translational machinery in higher eukaryotes, including ribosomal proteins, eEF1 and eEF2, and poly(A)‐binding proteins as well as many others, are regulated at the translational level. The structural hallmark common to all these mRNAs is the presence of a 50 ‐TOP tract, which contains a short polypyrimidine stretch (4–14 nucleotides) immediately adjacent to the 50 ‐cap (Meyuhas, 2000). Under conditions of nutrients or growth factor deprivation, or following the initiation of a differentiation program, the translation of 50 ‐TOP‐containing mRNAs is potently repressed. Conversely, stimulation by nutrients, hormones, cytokines, or growth factors overcomes translational repression. General components of the translational machinery as well as several trans‐acting factors have been proposed to modulate 50 ‐TOP activity (Meyuhas and Hornstein, 2000). Very early following mitogenic stimuli, ribosomal protein S6, whose phosphorylation is carried out in mammalian cells by two closely related kinases S6K1 and S6K2, becomes fully phosphorylated. All of the phosphorylation sites are conserved between the two proteins, but most of the studies are performed with S6K1, because S6K2 was discovered much later than S6K1. The fact that S6K1 activation precedes the translational activation of TOP mRNAs led to a model where rpS6 was the physiological target through which the S6 kinases mediated their effects on cell growth and TOP mRNAs translation (Jefferies et al., 1997; Fumagalli et al., 2000). Ribosomal protein S6 is a downstream target of the PI3K pathway, but the activation of S6K1 also hinges on the mTOR pathway (Caldarola et al., 2004). Very recent studies performed in diverse experimental models, including synaptoneurosome preparations or cultured hippocampal neurons, have shown that multiple forms of neuronal activity stimulate translational efficiency in association with the increased phosphorylation of rpS6, eIF4E, and 4E‐BP1. This phosphorylation was sensitive to rapamycin and also to U0126 (the specific inhibitors of mTOR and MAPKs pathways respectively) treatment, indicating the participation of mTOR/ERK signaling pathways. The authors suggest that stimulation of 50 ‐TOP‐dependent translation by rpS6 phosphorylation could play a key role during the establishment of long‐lasting forms of synaptic plasticity and memory (Kelleher et al., 2004a, b).


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More recent studies, however, question the role of rpS6 phosphorylation in the mechanism of 50 ‐ TOP mRNAs translation (Stolovich et al., 2002; Fingar et al., 2004; Pende et al., 2004); and other physiological targets of S6K1, which includes eIF4B, have been proposed (Hay et al., 2004; Raught et al., 2004).

3.2.2 Control at Internal Ribosome Entrance Internal translation initiation is a process initially described in picornavirus RNA whereby highly structured RNA elements directly recruit 40S subunits close to the initiator (AUG) codon on the mRNA and promote translation in the absence of a functional 50 ‐ cap‐binding protein complex eIF4F (Pelletier et al., 1988). Most of the IRES‐mediated translation, although a cap‐independent mechanism, requires most of the canonical translation factors, eIF3, eIF4A, eIF4B, at least the central domain of eIF4G, and Met‐tRNAimet for translation (Carter et al., 2000; Hellen et al., 2001). However, certain viral IRESs such as the cricket paralysis virus IRES occupy the ribosomal P site and assemble the 80S ribosomes even in the absence of any assisting initiation factor or Met‐tRNAi (Jan et al., 2003). Remarkably, the list of reported cellular mRNAs containing a regulated IRES element is growing daily. It is now accepted that this mechanism may be an adaptive cellular switch to maintain synthesis of key survival proteins under conditions of stress, cell cycle stage, or apoptosis, when cap‐dependent initiation is disfavored. In fact, their gene products are translated during the G2/M phase of the cell cycle, growth control, hypoxia, cellular stress (chaperons), apoptosis, and tissue‐specific translation (van der Velden et al., 1999; Belsham et al., 2000; Carter et al., 2000; Hellen et al., 2001; Meijer et al., 2002). Details of specific RNA structures and proteins involved in IRES function remain obscure. Recently, it has been reported that induction of cat‐1 (cationic amino acid transporter‐1) IRES in response to nutritional stress requires both phosphorylation of eIF2a and translation of a small uORF encoding a 48‐residue peptide. By using three different approaches the authors conclude that the IRES structure is dynamic and created by RNA–RNA interactions between the 50 ‐end of the leader and downstream sequences (Yaman et al., 2003). It has been demonstrated that N‐methyl‐D‐aspartate (NMDA)‐receptor‐mediated downregulation of translation induces the synthesis of specific proteins such as a‐Ca2þ/calmodulin‐dependent kinase II (a‐CaMKII) at the developing synapses (Scheetz et al., 2000). Because certain synaptically located mRNAs, including the a‐CaMKII mRNA, contain IRES elements (Pinkstaff et al., 2001), internal initiation may be at least one of the mechanisms used by the neurons to translate dendrite‐specific mRNAs under conditions where global cap‐dependent protein synthesis is inhibited.

3.2.3 Upstream Open Reading Frames in the 50 ‐UTR: Reinitiation In the majority of natural eukaryotic mRNAs, the start site for translation is the first AUG (sAUG) codon and their flanking sequences modulate the efficiency with which the sAUG codon is recognized as a stop signal during the scanning phase of initiation. When potentially active upstream initiation codons in the 50 ‐UTR region of an mRNA are present, three escape mechanisms, reinitiation, leaky scanning, and possibly internal initiation, allow access to sAUG codons, which although not the first, are still close to the 50 ‐end of the mRNA (Kozak, 1999). Reinitiation is a relatively rare event that is produced because after a ribosome translates the first uORF and reaches the termination codon, the 40S subunit may hold on to the mRNA, resume scanning, and reinitiate at a downstream AUG codon. In eukaryotes, reinitiation is most efficient when uORF terminates some distance before the start of the next cistron, but in general, the uORFs are usually inefficient and they are employed to decrease the initiation efficiency on the sAUG preceding the main ORF (Kozak, 2001; Meijer et al., 2002). Most interesting is the fact that this mechanism may also promote gene‐specific translational control (Kozak, 2001) and the first and best‐characterized example has emerged from studies in Saccharomyces cerevisiae (Hinnebusch, 1993). When the cells are starved of amino acids and glucose or subjected to other type of stress, general protein synthesis is repressed by phosphorylation of eIF2a by GCN2 kinase. However, the translation of transcription factor GCN4 mRNA, which is an

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activator of proteins that regulate amino acid biosynthesis, is specifically activated by a reinitiation‐ dependent mechanism (Dever et al., 1992; Hinnebusch, 1997; Meijer et al., 2002; Gebauer et al., 2004). The 50 ‐UTR of GCN4 mRNA contains four short ORFs upstream of the GCN4 initiation codon and each of them, especially uORF4, abolishes almost all mGCN4 translation. Under normal nutritional conditions, most of the small ribosomal subunits are capable of reinitiating upon reaching uAUG4. After translation of uORF4, ribosomes resume translation and leave the mRNA before reaching the GCN4 ORF. In the presence of elevated eIF2(aP) levels, ternary complex formation is compromised, and not all the ribosomal subunits are loaded in time for initiation at uORF4, instead, they are reloaded while scanning the remaining of the 50 ‐UTR and, hence, reach the GCN4 ORF (Meijer et al., 2002; Gebauer et al., 2004). In higher eukaryotes, very recent studies have demonstrated that under endoplasmic reticulum stress, general translation is inhibited upon eIF2a phosphorylation by PERK, while the translation of activating transcription factor 4 (ATF4) is specifically upregulated (Harding et al., 2002; Rutkowski et al., 2004). Mouse ATF4 50 ‐UTR contains two overlapping ORFs and although the mechanism of translational control of ATF4 is not identical with that of GCN4, both depend on uORFs and translational repression by increased eIF2(aP) (Meijer et al., 2002; Blais et al., 2004; Vattem et al., 2004). Reinitiation has been shown in only a small percentage of uORF‐containing messengers and in general is dependent of eIF2a phosphorylation. It can play a role in the synthesis of different protein isoforms encoded by the same mRNA or in tissue‐specific translation (Meijer et al., 2002). All together these results suggest that this mechanism of translation reinitiation is conserved from yeast to mammals and probably contributes to limit the translation of proteins including cytokines, transcription factors and other proteins, which could be harmful to the cell if overproduced.

3.2.4 Control by Cis‐Acting Elements Activated by Trans‐Acting Factors As already mentioned (see ‘‘Cis‐Acting Elements Dependent on Trans‐Acting Factors’’), the trans‐acting factors engaged in translational control are mainly RNA‐binding proteins that modulate the translation of specific mRNAs. We will describe in some more detail the mechanism of polyadenylation‐induced translation because it acquires importance particularly in the regulation of gene‐specific translation in dendrites in response to synaptic stimulation (Ritcher et al., 2002; Steward et al., 2003; Kelleher et al., 2004b). This extremely complex mechanism has been studied most extensively in Xenopus oocytes maturation (Hodgman et al., 2001; Mendez at al., 2001) where dormant mRNAs containing short poly(A) tails must be elongated for translational initiation to occur. Polyadenylation requires the participation of two cis‐acting elements, CPE and the poly(A) tail, the trans‐acting factor CPEB, the cytoplasmic form of cleavage and polyadenylation specificity factor (CPSF), and the poly(A) polymerase (PAP). After the induction of maturation, CPEB is phosphorylated by Aurora A (Eg2) kinase and recruits CPSF and PAP into an active polyadenylation complex (Hodgman et al., 2001; Sarkissian et al., 2004). In addition, CPE/CPEB interaction acts indirectly as a masking factor, but the main protein that mediates translational repression is maskin, another CPEB‐interacting factor (Stebbins‐Boaz et al., 1999). In oocytes, maskin not only binds CPEB but also eIF4E; this binding blocks eIF4E interaction with eIF4G and causes repression of translation (masking) prior to oocyte maturation. During oocyte maturation at a time corresponding with polyadenylation, the interaction between maskin and eIF4E is disrupted; eIF4E can interact with eIF4G and cap‐dependent translation is restored. It seems plausible that polyadenylation could be involved in the dissociation of maskin from eIF4E (Hodgman et al., 2001; Mendez et al., 2001). Compelling evidences support that this mechanism may regulate the translation of a‐CaMKII mRNA at synapses. Wu et al. (1998) demonstrated that NMDA receptor activation triggers polyadenylation of a‐CAMKII mRNA and expression of the protein. Besides, NMDA receptor signaling results in CPEB phosphorylation via ERK‐activated Aurora kinase and a‐CaMKII mRNA polyadenylation at synapses (Huang et al., 2002). A very recent work raises the hypothesis that phosphorylation of CPEB by CaMKII may promote protein synthesis from other CPE‐containing dendritic mRNAs during the induction of hippocampal LTP (Atkins et al., 2004).


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3.2.5 Control by MicroRNAs A new field in translational control has emerged with the view that not only protein factors through protein–RNA interactions, but also microRNA (miRNA)–RNA interactions are capable of repressing translation (Gebauer et al., 2004). Several hundreds of miRNAs have recently be cloned from a wide range of organisms across phylogeny and their functions in mammals are yet to be defined (Krichevsky et al., 2003). In neurons, a dendritic nontranslatable small RNA, BC1RNA, which does not contain a protein‐coding sequence, acts as a repressor in both cap‐dependent and internal entry modes, and a functional role of BC1RNA as a mediator of translational control in local protein synthesis has been suggested. This mechanism, however, is distinct from that of miRNAs because BC1RNA specifically inhibits 48S formation and the interaction with eIF4A and with poly(A) seems to be required (Wang et al., 2002). The fragile X mental retardation protein (FMRP) is an RNA‐binding protein that is associated to polysomes both in the cell body and dendrites of neurons (Jin et al., 2003; Jin et al., 2004). Loss of FMRP, which is the defect responsible for fragile X syndrome in humans, increases long‐term depression in mouse hippocampus and growing evidences support the role of FMRP as a repressor of translation of specific mRNAs in dendrites (Bear et al., 2004). FMRP may repress translation of specific mRNAs by several different ways including a direct interaction in the nucleus with the specific target mRNA (hairpin structure) and proteins to form messenger ribonucleoprotein (mRNP) complex. This complex is exported to the cytoplasm where it can be associated to ribosomes and regulate protein synthesis in the cell body of the neuron either directly or after interaction with the RNA‐induced silencing complex or the miRNA pathway. Alternatively, FMRP complex can be transported into the dendrites to regulate local proteins synthesis (Jin et al., 2004). In fact, it has been reported that FMRP may indirectly bind to dendritic‐specific mRNAs through its association to noncoding RNAs such as BC1RNA (Zalfa et al., 2003).

4

Cerebral Ischemia‐Induced Downregulation of Translation

4.1 Introduction One of the first cellular functions to be affected by cerebral ischemia is protein synthesis. In rodents, downregulation of brain protein synthesis is observed when cerebral blood flow is only 70–80% of its normal rate, a value that is even higher than the perfusion rate required to maintain ATP production, suggesting that factors other than energy supply are involved in translation downregulation. Upon reoxygenation, protein synthesis is strongly inhibited and its recovery is much slower than that of energy metabolism, the recovery being dependent on the ischemic period duration (Xie et al., 1989; Hossmann, 1993; Iadecola, 1999). Moreover, a persistent protein synthesis inhibition in selectively vulnerable neuronal cells preceding neuronal death has been observed (Dienel et al., 1980; Thilmann et al., 1986; Hossmann, 1993; Krause et al., 1993). The striking correlation between neuronal vulnerability and downregulation of protein synthesis suggests that this cellular process is a critical part in the cascade of pathogenetic events leading to ischemic cell death. Investigations in the last 10 years have focused on the study of the mechanisms involved in ischemia‐induced translation inhibition as a source of clues to find out the potential role of protein synthesis in the manifestation of selective brain injury.

4.2 Experimental Models of Cerebral Ischemia Kleihues and Hossmann, between 1971 and 1975 (Hossmann, 1993), carried out the first investigations to study the effect of brain ischemia on protein synthesis using a model of prolonged cerebrocirculatory arrest in cats and monkeys. A considerable number of papers published between the 1970–1990s, with different in vivo experimental models of ischemia in other animal species such as rats and gerbils, have

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confirmed the ischemia‐induced downregulation of translation. All these studies include transient global ischemia by cardiac arrest, two‐vessel and four‐vessel occlusion models, and permanent or transient focal ischemia performed by middle cerebral artery occlusion (Cooper et al., 1977; Morimoto et al., 1978; Dienel et al., 1980; Nowak Jr et al., 1985; Xie et al., 1989; Araki et al., 1990; Widmann et al., 1991; Krause et al., 1993; Hata et al., 1998). Ischemia‐induced inhibition of translation has also been studied with cell‐free preparations from ischemic rat brain as well (Erdogdu et al., 1993; DeGracia et al., 1993; Burda et al., 1998). Protein synthesis is also inhibited in neuronal cultured cells or slices subjected to glucose deprivation and anoxia, indicating that these in vitro models may be useful and inexpensive tools to study the mechanisms implicated in ischemia‐induced downregulation of translation (Ouellette et al., 1983; Raley‐ Susman et al., 1990a, b; Raley‐Susman et al., 1995; Munõz et al., 2000). Even though they have several drawbacks, the most obvious being that they lack real ischemia, i.e., reduced blood flow (Choi, 1990).

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Regulation of Protein Synthesis During Cerebral Ischemia

5.1 Ischemic Period Our discussion will be mostly focused in reviewing the latest results concerning translation regulation under transient global ischemia, in an effort to clear up the role this regulatory mechanism may play in delayed ischemic damage. It is generally believed that during ischemia protein synthesis is completely suppressed because energy‐rich phosphates required for the acylation of tRNA and the formation of peptide bonds are rapidly depleted. Polyribosomes remain intact because the three steps of translation come to a halt—‘‘ischemic freeze’’ (Hossmann, 1993). While these statements are true, results from several laboratories have showed that several components of the translational machinery are modified immediately after the ischemia period (Neumar et al., 1998; Martıń de la Vega et al., 2001a; Mengesdorf et al., 2002; Garcıá et al., 2004b). We induced incomplete forebrain ischemia (30 min) in rats by using the four‐vessel occlusion model, and the levels and the phosphorylation status of different initiation factors and protein kinases were determined in extracts from the cortex and the hippocampus, as representative of resistant and vulnerable brain regions to ischemia, respectively. We found that 4E‐BP1 becomes fully dephosphorylated during the ischemia period and as a result we must expect the disposability of eIF4E compromised after this period (> Figure 1-3) (Martıń de la Vega et al., 2001a; Garcıá et al., 2004b). The same result was obtained with an in vitro model of ischemia using NGF‐differentiated PC12 cells exposed to glucose deprivation and anoxia (Martıń et al., 2000b). In addition, eIF4E and eIF4G protein levels, independently of the method used to induce ischemia, were lower after the ischemic period (> Figure 1-3), this result being probably due to the proteolysis caused by calcium‐mediated calpain activation (DeGracia et al., 1996; Neumar et al., 1995, 1996, 1998; Burda et al., 1998; Martıń de la Vega et al., 2001a; Mengesdorf et al., 2002; Garcıá et al., 2004b). The levels of phosphorylated eIF4E and eIF4G also decrease during the ischemic period, while the levels of phosphorylated eIF2a were similar to those of the controls (Martıń de la Vega et al., 2001a; Garcıá et al., 2004b). A complete dephosphorylation of eIF2Be after transient focal ischemia has also been reported (Mengesdorf et al., 2002). Most of the kinases responsible for the phosphorylation of initiation factors, including Akt/PKB, ERK1/2 and S6K1, were dephosphorylated, i.e., inactivated during ischemia. (Martıń de la Vega et al., 2001a; Mengesdorf et al., 2002; Garcıá et al., 2004b). In the presence of a normal energetic supply, this situation would imply an inhibition of cap‐dependent and 50 ‐TOP mRNAs translation. However, the translation of strong mRNAs with low secondary structure in the 50 ‐UTR region such as those of heat‐shock proteins or mRNAs containing IRES could be upregulated. Further studies will be necessary to know whether or not the alterations in the translational machinery, induced during the ischemia period, have functional consequences in the strong inhibition of translation that occurs immediately after the onset of reperfusion. Nevertheless, this potential translational control could be, at least partially, responsible for the multiple penumbras that in molecular terms i.e., gene and protein expression, have been described in focal ischemia (Sharp et al., 2000).


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. Figure 1-3 Translational machinery during transient global ischemia. Protein synthesis is an energy requiring process, and consequently, completely suppressed during the ischemia period after transient global ischemia because of generalized energy failure. It is believed that polysomes remain intact because the three steps of protein synthesis come to a halt—‘‘ischemic freeze.’’ However, decreased levels of eIF4G, eIF4E, and eIF2B (striped lines), possibly caused by a calpain‐mediated proteolysis, have been observed. In addition, eIF2a as well as other initiation factors and protein kinases are completely dephosphorylated after this period

5.2 Early Reperfusion Period 5.2.1 Initiation Factors The fact that protein synthesis inhibition during reperfusion is associated to polysome disaggregation was indicative of a translational control mainly exerted at the rate‐limiting initiation step (Cooper et al., 1977; Burda et al., 1980; Hossmann, 1993). A potential regulation at the initiation step was further supported by the fact that eIF2B activity was found to be inhibited during ischemic reperfusion (Hu et al., 1993). With the model of transient global ischemia in the rat, we first demonstrated that in brain extracts the phosphorylation of eIF2a paralleled protein synthesis inhibition and ternary complex formation, besides being responsible for eIF2B activity inhibition (Burda et al., 1994). This phosphorylation occurred similarly in the cortex and the hippocampus (Burda et al., 1994; Martıń de la Vega et al., 2001a, b) (> Figure 1-4). The immunohistochemistry results showed in > Figure 1-4 supports Western blot results and definitively proves that eIF2(aP) levels return to control values after 4–6 h of reperfusion in the cortex as well as in the hippocampal subregions, although dephosphorylation of eIF2(aP) is visibly delayed in CA1 in comparison to other hippocampal subregions (> Figure 1-4b). Results obtained with an in vitro model of ischemia using NGF‐differentiated PC12 cells exposed to glucose deprivation and anoxia indicated that both the increase in eIF2(aP) levels and protein synthesis inhibition were induced mostly during the ischemic instead of the recirculation period (Munõz et al., 2000). While different mechanisms occurring in in vivo and in vitro models are unlike, a plausible explanation to this discrepancy could be that ATP levels are completely depleted after ischemia uniquely in the in vivo model. According to this hypothesis, eIF2a is not phosphorylated during ischemia in the in vivo model because the eIF2a kinase, as it occurs with all the other kinases studied, is completely dephosphorylated (inactive) within this period.

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. Figure 1-4 Phosphorylation of eIF2a during ischemic reperfusion. Panel a shows that eIF2a phosphorylation, as determined by isoelectric focusing slab electrophoresis followed by immunoblot assay using monoclonal antibodies against total eIF2a, is induced very early during reperfusion following transient global ischemia (30 min), but recovers to normal levels after 4–6 h of reperfusion. No differences were observed between the cortex and the hippocampus (Martıń de la Vega et al., 2001a, b). The immunohistochemistry experiments showed in Panel b are performed with antibodies against eIF2(aP) and confirm that eIF2a phosphorylation is triggered in resistant and vulnerable hippocampal subregions early at reperfusion. Although eIF2(aP) dephosphorylation is delayed in CA1 subregion, the factor is completely dephosphorylated after 4 h in all hippocampal subregions (Salinas and Burda unpublished results)

Increased eIF2a phosphorylation immediately after the beginning of reperfusion has been found in all the in vivo experimental models of global or focal ischemia studied so far (DeGracia et al., 1996; Burda et al., 1998; Althausen et al., 2001; Mengesdorf et al., 2002; Hayashi et al., 2003a). Thus, it is now well established that cytoplasmic eIF2a phosphorylation is the major site of early postischemic translational regulation. Although we could not detect any increased eIF2a kinase activity in our model of ischemia, studies from other laboratories performed with different in vivo ischemia models have revealed paralleled increases in PERK and eIF2a phosphorylation (Kumar et al., 2001; Hayashi et al., 2003a, b). It has been concluded that PERK is activated because the phosphorylated form of PERK is responsible for eIF2a phosphorylation in other stress situations (Harding et al., 1999, 2002). PERK activation does not exclude a potential modulation of eIF2(aP) levels by protein phosphatase 1 (PP1), which is the physiological phosphatase suggested to be responsible for eIF2a dephosphorylation in vivo (Novoa et al., 2001; Brush et al., 2003). Because the catalytic subunit of PP1 (PP1c) may be regulated by oxidation (O’Loghlen et al., 2003), an inactivation of PP1c caused by reperfusion‐induced oxidative stress (Chan, 1996; Lipton, 1999; Hou et al., 2002) could collaborate to increase eIF2(aP) levels during reperfusion (Munõz et al., 2000; Martıń de la Vega et al., 2001b).


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A decrease in the formation of eIF4F complex and diminished eIF4E, eIF4G, and eIF2Be protein levels have been observed during early reperfusion that could act concomitant to eIF2a phosphorylation in inhibiting overall translation (Martıń de la Vega et al., 2001a; Mengesdorf et al., 2002).

5.2.2 Endoplasmic Reticulum Dysfunction The ischemia‐induced accumulation of protein aggregates (Hu et al., 2000), activation of PERK, and phosphorylation of eIF2a, are hallmarks indicative that ischemia may be one of the pathophysiological conditions that triggers ER stress (Paschen and Doutheil, 1999). It has been demonstrated that this condition in turn induces a coordinated transcriptional and translational adaptive program of response to stress called the unfolded protein response (UPR) (Harding et al., 2002; Kaufman, 2002; Rutkowski et al., 2004). Efforts have been made in the last 5 years to find the correlation between UPR dysfunction and ischemic death. While these studies showed increases in the mRNAs of several components of the UPR, including transcription factors XBP‐1, ATF4, and ATF6, the glucose‐regulated protein of 78 kDa (GRP78/ Bip) and the growth arrest and DNA damage protein 34 (GADD34), they failed to find increased expression of the corresponding protein products (Doutheil et al., 1999; Kumar et al., 2003; Paschen et al., 2003). Two of these studies reached the conclusion that the strong inhibition of translation in the early period after transient brain ischemia critically limits new synthesis of all the proteins and blocks an effective UPR against ER stress. However, the model of focal ischemia used in one of the studies (Paschen et al., 2003) and the short reperfusion times studied with the model of cardiac arrest (Kumar et al., 2003) do not allow to establish any correlation between ER stress and delayed CA1 hippocampal death. Hayashi et al. (2003b) used longer reperfusion times after transient global ischemia and showed an increased expression of GRP78 from 4 h to 2 days after reperfusion at the hippocampal CA1 cell layer. Interestingly, GRP78 levels increase after ischemic preconditioning. Although parallel studies at resistant brain areas are not described, the authors suggest that the increased GRP78 expression induced by preconditioning may reduce ER stress and eventually delay neuronal death after ischemia (Hayashi et al., 2003b). In our laboratory we have not observed any change in GRP78 levels at 4 h of reperfusion with a similar model of ischemia; instead, we found an increase in GADD34 expression both in the cortex and the hippocampus at this reperfusion time (Garcıá et al., 2004b). Increased GADD34 levels in the cortex, but not in the CA1 subfield of superoxide dismutase‐1 overexpressing rats, after 1–6 h of transient global ischemia has also been reported very recently (Paschen et al., 2004). In conclusion, it has been proposed that the mechanism of eIF2a phosphorylation constitutes a classical stress‐induced acute response, common to vulnerable and nonvulnerable cells, which if persisting, can have deleterious effects on cell survival (Martıń de la Vega et al., 2001a; Paschen, 2003; Garcıá et al., 2004b; DeGracia, 2004). However, there are no compelling evidences for the moment that support a specific and persistent ER stress caused by eIF2a phosphorylation being responsible for the delayed neuronal death observed in ischemia‐vulnerable brain regions.

5.3 Late Reperfusion Period Polysome profile determination, as well as protein synthesis rate measurements determined in both in vivo and in vitro methods, indicate that protein synthesis inhibition, although partially recovered, is still substantial after 6 h reperfusion in resistant areas, while remaining inhibited in hippocampal CA1 subregion (Thilmann et al., 1986; Hossmann, 1993; Martıń de la Vega et al., 2001a; Burda et al., 2003).

5.3.1 Initiation Factors and Protein Kinases Most of the components of the translational machinery, 4E‐BP1, eIF4E, eEF2, and S6K1, which were completely or partially dephosphorylated after the ischemic period, progressively recovered their

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phosphorylated status with increasing reperfusion time (Althausen et al., 2001; Martıń de la Vega et al., 2001a; Mengesdorf et al., 2002; Garcıá et al., 2004b). eIF2B activity exactly paralleled eIF2a phosphorylation, decreasing at early reperfusion but reaching control values at late reperfusion (Martıń de la Vega et al., 2001a; Garcıá et al., 2004b). Since GSK3 is dephosphorylated (activated) during ischemia but phosphorylated (inactivated) at early reperfusion (Salinas and Burda; unpublished results), regulation of eIF2B activity through this kinase should not be expected. A long‐lasting dephosphorylation of eIF2Be starting during ischemia in a model of transient focal ischemia has also been reported (Mengesdorf et al., 2002). The phosphorylated levels of eIF4G, which strictly paralleled those of Akt were significantly higher after 4–6 h reperfusion than in control, but as already discussed, the potential role that eIF4G phosphorylation may play in translational regulation is at present time unknown (Garcıá et al., 2004b).

5.3.2 eIF4G and Calpain Activation eIF4G levels that were lower than controls during ischemia, did not normalize during reperfusion and could be responsible, at least in part, of the persistent translation inhibition observed at late reperfusion times (> Figure 1-5). Again results with the in vivo model are in disagreement with those obtained with the in vitro model of ischemia, where eIF4E and eIF4G levels do not change either during ischemia or recirculation (Martıń et al., 2000b). Thus, as far as the regulation of eIF2 phosphorylation and eIF4E and eIF4G levels are concerned, the discrepancies observed between the in vivo and the in vitro models question the efficacy of the later for the study of transient cerebral ischemia. Caspase‐3 and calpain are cysteine proteases with a broad spectrum of substrates, which are activated following transient global and focal ischemia in a complex spatiotemporal pattern. Caspase and calpain inhibitors block not only the proteolytic activities of the proteases, but also the cellular process of death itself in vulnerable neurons in a synergistic manner. This result supports its role in protecting neurons, suggesting cross talk between caspase and calpain activities (Li et al., 1998; Rami et al., 2000; Zhang et al., 2002; Rami, 2003). eIF4G is a substrate of both proteases, but calpain seems to be responsible for the maximal cleavage of eIF4G during ischemia and reperfusion (Neumar et al., 1996, 1998; Garcıá et al., 2004b). Calpain activation is modulated by calpastatin, an endogenous inhibitor of calpain, which is present in lower concentrations in vulnerable pyramidal cells than in resistant CA3 cells. Moreover, calpastatin is degraded if the calpain/calpastatin ratio increases (Fukuda et al., 1990; Rami, 2003). Given that we have found increased calpain activity during reperfusion in both the cortex and the hippocampus, a lower calpain/calpastatin ratio in the later could explain the lower eIF4G levels found in the CA1 hippocampal subfield (Martıń de la Vega et al., 2001a; Garcıá et al., 2004b). All together, the results accumulated until now suggest that the ischemia‐induced translational repression is probably mediated by the summation of multiple altered regulatory sites that are selectively impaired with the progression of ischemia and reperfusion periods. Given the complexity of the translational machinery, those mechanisms already studied, as well as others implying translation factors such as eIF1, eIF3, eIF5, or eIF5B could be functionally important. Ischemia is one of the strongest stimuli of gene induction in the brain (Nowak et al., 1999; Hou et al., 2002). With the aid of microarray analysis, it has been confirmed that many different gene systems related to reperfusion processes of brain injury, repair, and recovery are up‐ or downregulated (Jin et al., 2001; Read et al., 2001; Lu et al., 2003; Gilbert et al., 2003). As proposed in > Figure 1-5, translation inhibition induced by the clear‐cut rise in eIF2(aP) levels at early reperfusion, its moderate slow dephosphorylation, and the long‐lasting recovery of global translation during late reperfusion caused by eIF4G degradation and possibly by other factors, represents a liable situation to induce a decrease in cap‐dependent translation and upregulation of inefficient mRNAs’ translation (uORFs, structured 50 ‐UTR, IRES, 50 ‐TOP), which in turn may definitively help the induction of a survival program in resistant neurons. We still do not know because translation repression fails to recover in vulnerable cells during late reperfusion, either those inhibitory mechanisms common to vulnerable and resistant cells remain operative only in the former, or alternatively, other inhibitory mechanisms are induced exclusively in vulnerable cells at late reperfusion.


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. Figure 1-5 Translation regulation during reperfusion following transient global ischemia in resistant neurons. Multiple altered mechanisms regulate translation following ischemic reperfusion time (see > Sect. 5). At very early reperfusion (10 min), PERK activation and possibly PP1 inhibition synergistically promote eIF2a phosphorylation. As a consequence, eIF2B is sequestered, ternary complex formation is inhibited, and translation is strongly repressed. Following reperfusion times, the equilibrium of the phosphorylation reaction is displaced to the dephosphorylated protein, because PERK phosphorylation decreases and PP1 is activated, hence, eIF2(aP) levels slowly decrease and translational rate increases. Although eIF2(aP) levels reach control values after 4–6 h reperfusion, translation remains inhibited, very probably due to the lower levels of eIF4G as well as other unknown mechanisms. As described in the text, this situation may be compatible with cap‐dependent translation inhibition and upregulation of structured mRNAs, which in turn may help the induction of a survival program for resistant neurons

5.4 Translation Control in Delayed Neuronal Death There are changes in intrinsic basal levels of variables that are different in vulnerable (CA1 hippocampal pyramidal cells) and resistant cells that might reasonably account for vulnerability of CA1 to global ischemia. These include among many others, decreased slow recovery of pH, zinc accumulation, maintained elevation of free fatty acids, progressive loss of mRNAs, decreased superoxide dismutase, increased caspase mRNA levels, lower calpastatin basal levels, delayed decrease in GluR2/GluR1 receptor ratio, and permanently repressed translation (Maruno et al., 1990; Lipton, 1999). The lower global translational rate and the delayed eIF2(aP) dephosphorylation of vulnerable neurons might modify the equilibrium between survival and death programs, turning the survival program into a death program. Besides, the levels of short half‐life cellular proteins would be most affected in vulnerable cells (> Figure 1-6). MacManus et al. (2004) have used a very accurate approach to examine gene expression after ischemia, which consists in analyzing

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. Figure 1-6 Potential mechanisms of translational control in the processes of ischemic‐delayed neuronal death and tolerance acquisition

by microarray those mRNAs bound to polyribosomes instead of the total transcript. One of their conclusions is that the presence of IRES or 50 ‐TOP mRNAs does not appear to be the basis of selection for translation in the ischemic brain. However, this conclusion does not contradict our hypothesis, because the changes in gene expression induced in a 20‐h‐reperfused brain hemisphere after 1 h focal ischemia are very possibly quantitatively and qualitatively different to those triggered in a vulnerable region after global postischemic reperfusion. In any case this method, ‘‘translation state analysis,’’ or the arrays of proteins will definitively help to delineate the pattern of proteins with significant role within the ischemic process.

5.4.1 Control of Protein Expression: GluR2 According to the secondary reperfusion injury hypothesis for delayed neuronal death, a persistent rise in intracellular [Ca2þ], which occurs specifically in hippocampal CA1 neurons between 1 and 3 days after induction of global ischemia, triggers a secondary failure of many cellular functions leading to delayed cell death (Kristiań et al., 1998; Siesjo¨ et al., 1999). It has been demonstrated that GluR2, the subunit that limits Ca2þ permeability of alpha‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazolenepropionic acid (AMPA)‐type glutamate receptor, is specifically and acutely reduced in CA1 neurons after transient forebrain ischemia (Gorter et al., 1997). In addition, different studies have provided evidences for the contribution of Ca2þ‐permeable AMPA receptors to the pathogenesis of ischemia‐induced neuronal death (Tanaka et al., 2000; Liu et al., 2004). Transcriptional mechanisms that implie either the transcription factor cAMP response element‐ binding protein (CREB) or the gene silencing transcription factor neuronal repressor element‐1 (REST) have been proposed to regulate GluR2 subunit expression after transient global ischemia (Calderon et al., 2003; Liu et al., 2004). The presence of multiple transcription sites for the GluR2 gene results in a heterogeneous population of GluR2 transcripts in vivo that differ in the length of their 50 ‐UTRs. Interestingly, the


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relative amount of GluR2 transcripts with long versus short 50 ‐UTRs varies across brain regions and the subpopulation of GluR2 mRNAs with long 50 ‐UTRs are subject to translation inhibition through a polymorphic repeat motif predicted to form a secondary structure in vivo (Myers et al., 2004). Hence, the potential role of translational control in the downregulation of GluR2 expression in vulnerable neurons following transient global ischemia deserves to be fully addressed.

5.4.2 Inhibition by Calpain Activation Elevation of free intracellular [Ca2þ] induces calpain activation (Rami et al., 2000; Zhang et al., 2002; Rami, 2003). It has been postulated that calpain cleaves among others the p35 neural‐specific activator of cyclin‐ dependent kinase 5 (Cdk5) into its truncated form p25 (Tsai et al., 1994; Patrick et al., 1999; Lee et al., 2000). The accumulation of p25 induces a prolonged activation of Cdk5, which phosphorylates the NR2A subunit of the NMDA receptor, enhancing its function and promoting hippocampal CA1 delayed cell death (Wang et al., 2003; Shelton et al., 2004). Downregulation of protein synthesis at reperfusion periods longer than 6 h has been studied by in situ autoradiography analysis of amino acid incorporation, but data about the activity of the individual components of the translational machinery are not available yet. Without excluding the participation of other deleterious processes induced by the elevation of free intracellular [Ca2þ] (Lipton 1999; Kristiań et al., 1998; Brostrom et al., 2003; Rami, 2003), a further repression of translation caused by calpain‐induced eIF4G cleavage would not be unexpected, and could absolutely collaborate to the delayed neuronal death (> Figure 1-6). Besides, given the broad spectrum of calpain substrates, the cleavage of other translational components should not be discarded.

5.5 Translational Control in Ischemic Preconditioning A brief ischemic episode, which is in itself nonlethal (ischemic preconditioning), confers tolerance to subsequent ischemic insults (Kitagawa et al., 1990; Barone et al., 1998). Results based on autoradiography and electron microscopy first showed that the ischemic tolerance (IT) is closely related to the facilitated recovery from suppressed protein synthesis in the brain after ischemia (Furuta et al., 1993; Kato et al., 1995).

5.5.1 Improvement of Initiation at Early Reperfusion Our recent results suggest that two different mechanisms are essential for the acquisition of ischemic tolerance, at least in the CA1 sector of hippocampus. The first mechanism implies a significant reduction in translation inhibition after lethal ischemia, especially at an early time of reperfusion, in both vulnerable and nonvulnerable neurons. For the acquisition of full tolerance, a second mechanism, highly dependent on the time interval between preconditioning and lethal ischemia, is absolutely necessary (Burda et al., 2003). Ischemic preconditioning induces a complete reorganization of the gene program of ischemic neurons and it has been hypothesized that the ischemic tolerance represents the acquisition by vulnerable cells of properties shown by less vulnerable cells (Lipton, 1999). As it occurs with heat shock protein of 72 kDa expression, the second mechanism indicated above might involve the transcriptional induction and corresponding synthesis of the protective proteins that prevent the delayed death of vulnerable neurons after lethal ischemia (Burda et al., 2003). In our model of preconditioning, the partial recovery in the translational rate induced at early reperfusion after lethal ischemia, which is sufficient to induce protection, is induced in the cortex as well as in the hippocampus. This result may suggest that vulnerable cells have acquired the properties of resistant cells, thus, translation slowly recovers its normal rate during late reperfusion in both regions (> Figure 1-6). Alternatively, the possibility that preconditioning may eliminate the potential inhibitory mechanisms specifically developed in vulnerable cells during late reperfusion should not be excluded.

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Ischemic preconditioning does not prevent the transient phosphorylation of eIF2a, which occurs at 30 min reperfusion. eIF2(aP) levels return to control values within 4 h both in the cortex and hippocampus (Garcıá et al., 2004b). Nevertheless, we do not know whether or not the dephosphorylation of the factor at this period of time occurs faster in rats with acquired tolerance. Therefore, the exact mechanisms implicated in the preconditioning‐induced attenuation of translation remains unknown, but a helpful effect due to a better preservation of the translational machinery during the ischemia period, as suggested by the observed increase in 4E‐BP1 phosphorylation (Garcıá et al., 2004b), should not be discarded.

5.5.2 Control of Survival Proteins: Expression of GADD34 Very interestingly, the partial preconditioning‐induced recovery of translation is enough to induce an early onset of GADD34 expression—after 30 minutes instead of 4 h. As recently proposed (Novoa et al., 2001; Kojima et al., 2003), the increased levels of this protein may accelerate the programmed recovery of translational repression, and consequently, abolishes ER stress whose dysfunction is detrimental to neurons (Kumar et al., 2003; Paschen, 2003). Moreover, as is the case with GADD34 and GRP78 (Hayashi et al., 2004), the synthesis of proteins engaged in cell survival programs, whose genes are upregulated by tolerance (Dhodda et al., 2004; Kawahara et al., 2004), may be facilitated.

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Conclusion

Whereas considerable progress has been made in understanding the mechanisms implicated in ischemia‐ induced translation repression, the extraordinary complexity of the translational machinery makes it a difficult task. Whether or not the impairment of the translational machinery is the cause of any form of ischemic neuronal death remains to be firmly clarified. However, what is clear is that translational control plays a key role in several steps of the development and prevention of ischemic injury. Any progression in the knowledge of the mechanisms implicated will provide potential targets for future therapeutic strategies directed to prevent pathophysiological events associated within the ischemic brain.

Acknowledgments We apologize to investigators whose work was not cited or discussed in detail due to space limitations. We want to thank our collaborators Alberto Alca´zar, Juan L. Fando, M. Elena Martıń, and Victor M. Gonza´lez for valuable suggestions and the critical reading of this chapter. We are also indebted to our fellows, Milina Hrehorovska´, Lidia Garcıá, Cristina Martıń de la Vega, Francisco Munõz, Celia Quevedo, and Ana O’Loghlen, for the excellent work performed in the last years. Work from the authors’ own laboratory was supported by grants FIS 02/0304 and 05/0312 (Ministerio de Sanidad y Consumo) and SK‐VEGA 2/ 3219/23 (Slovak Academy of Sciences).

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2

Cell Adhesion Molecules of the Immunoglobulin Superfamily in the Nervous System

P. S. Walmod . M. V. Pedersen . V. Berezin . E. Bock

1 1.1 1.2 1.3 1.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 Defining an Adhesion Molecule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 Restrictions in Protein Types and Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 Cis‐Interactions, Trans‐Interactions, and Homophilic and Heterophilic Interactions . . . . . . . . . . . 44 The Immunoglobulin Superfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44

2 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8

Myelin Protein Zero . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.7.1 3.7.2 3.8

Integrin‐Associated Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52

4 4.1 4.2 4.2.1 4.2.2 4.2.3 4.2.4 4.2.5 4.2.6 4.2.7 4.3 4.3.1

The CTX Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 CXADR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 54 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57 IGSF11 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58

#


36

2

Cell adhesion molecules of the immunoglobulin superfamily in the nervous system

4.3.2 4.3.3 4.3.4 4.3.5 4.3.6 4.3.7

Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59

5 5.1 5.2 5.3 5.4 5.5 5.6 5.7

Neuroplastin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Isoforms and Primary Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60

6 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.7.1 6.7.2 6.8 6.9

Signal‐Regulatory Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65

7 7.1 7.2 7.3 7.4 7.5 7.6 7.7 7.7.1 7.7.2 7.8 7.9

Nectins (Poliovirus Receptor‐Related Proteins) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68 Nectin–Nectin Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71

8 8.1 8.2 8.3 8.4 8.5 8.6 8.7 8.7.1 8.7.2

Nectin‐Like Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 Necl–Necl Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76


2

8.8 8.9

Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

9 9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8 9.9

The IgLON Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 Interactions Within the IgLON Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

10 10.1 10.2 10.3 10.4 10.5 10.6 10.7 10.7.1 10.8

Activated Leukocyte‐Cell Adhesion Molecule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

11 11.1 11.2 11.3 11.4 11.5 11.6 11.7 11.7.1 11.7.2 11.8

Melanoma Cell Adhesion Molecule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85

12 12.1 12.2 12.3 12.4 12.5 12.6 12.7 12.8 12.9

Myelin‐Associated Glycoprotein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88

13 13.1 13.2 13.3

Neural Cell Adhesion Molecule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

37

38

2


13.4 13.5 13.6 13.7 13.7.1 13.7.2 13.8 13.9

Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94

14 14.1 14.2 14.3 14.4 14.5 14.6 14.7 14.8

Neural Cell Adhesion Molecule 2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97

15 15.1 15.2 15.3 15.4 15.5 15.6 15.7 15.7.1 15.7.2 15.8 15.9

The Robo Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 Isoforms and Primary Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Extracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100

16 16.1 16.2 16.2.1 16.2.2 16.2.3 16.2.4 16.2.5 16.2.6 16.2.7 16.2.8 16.3 16.3.1 16.3.2 16.3.3 16.3.4 16.3.5 16.3.6

The Contactin Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 F3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 TAG‐1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104


2

16.3.7 16.3.8 16.4 16.5 16.6 16.7

Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Contactin‐3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Contactin‐4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Contactin‐5 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Contactin‐6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105

17 17.1 17.2 17.2.1 17.2.2 17.2.3 17.2.4 17.2.5 17.2.6 17.2.7 17.2.8 17.3 17.3.1 17.3.2 17.3.3 17.3.4 17.3.5 17.3.6 17.3.7 17.4 17.4.1 17.4.2 17.4.3 17.4.4 17.4.5 17.4.6 17.5 17.5.1 17.5.2 17.5.3 17.5.4 17.5.5 17.5.6 17.5.7 17.5.8

The L1 Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 L1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 NrCAM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Neurofascin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112 CHL1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113

18

Down Syndrome Cell Adhesion Molecule and Down Syndrome Cell Adhesion Molecule‐Like 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115

18.1 18.2 18.3 18.4 18.5

39

40

2


18.6 18.7 18.7.1 18.8 18.9

Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Heterophilic Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Intracellular Binding Partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116

19 19.1 19.2 19.3 19.4 19.5

Sidekick 1 and 2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Isoforms and Protein Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 Homophilic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118

20 20.1 20.2 20.3 20.4

Putative Neural Adhesion Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 IGSF9 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 Punc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Nope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 TMEM25 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120


2

Abstract: Cell adhesion molecules (CAMs) are proteins mediating cell-cell or cell-extracellular matrix (ECM) interactions. CAMs are traditionally divided into four groups, the cadherins, the selectins, the integrins and CAMs belonging to the immunoglobulin superfamily (IgSF). The present chapter describes CAMs belonging to IgSF, that exclusively or in part, are expressed in the nervous system. The chapter includes descriptions of myelin protein zero (P0), integrin-associated protein (CD47), neuroplastin, activated leukocyte-cell adhesion molecule (ALCAM), melanoma cell adhesion molecule (MCAM), myelinassociated glycoprotein (MAG), the neural cell adhesion molecules 1 and 2 (NCAM, NCAM2), Down Syndrome cell adhesion molecule (DSCAM) and Down Syndrome cell adhesion molecule-like-1 (DSCAML1), sidekick 1 and 2 (SDK1, SDK2), signal-regulatory proteins (SIRPs), nectins, nectin-like proteins (necls), and members of the CTX, IgLON, Robo, contactin, and L1 families. The individual descriptions include sections describing the expression, isoforms, structure, posttranslational modifications, homo- and heterophilic binding partners, signaling and functions of the individual CAMs. The chapter demonstrates CAMs to be more than simple regulators of adhesion. Many CAMs are important mediators of intracellular signal transduction, and CAMs are involved in many biological phenomena including migration, proliferation, and differentiation of cells, as well as axonal guidance, neurite outgrowth, and synaptic plasticity and maturation. List of Abbreviations: 2‐AG, 2‐arachidonylglycerol; AA, arachidonic acid; BHV, bovine herpesvirus; BIT, brain immunoglobulin‐like molecule with tyrosine‐based activation motifs; CNS, central nervous system; ConA, concanavalin A; CRIB, Cdc42/Rac1 interactive binding; CSPG, chondroitin sulfate proteoglycan; DAG, diacylglycerol; DS‐CHD, Down syndrome congenital heart disease; E(number), embryonic day (number); ECM, extracellular matrix; EC module, extracellular module; F3, fibronectin type III; GAP, GTPase‐activating protein; GNA, Galanthus nivalis agglutinin; GPI, glycosylphosphatidylinositol; HGPPS, horizontal gaze palsy with progressive scoliosis; HIVAN, HIV‐associated nephropathy; HSPG, heparan sulfate proteoglycan; HSV, herpes simplex virus; Ig, immunoglobulin; IP3, inositol‐1,4,5‐trisphosphate; ITIM, immunoreceptor tyrosine‐based inhibitory motif; kb, kilobase; LPA, lysophosphatidic acid; LTD, long‐term depression; LTP, long‐term potentiation; Mbp, megabase pairs; MSD1, muscle‐specific domain 1; NMDA, N‐methyl‐D‐aspartate; NMR, nuclear magnetic resonance; NRSE, neuron‐restrictive silencer element; P(number), postnatal day (number); PIP2, phosphatidylinositol‐4,5‐bisphosphate; PI‐PLC, phosphoinositide‐specific phospholipase C; PNS, peripheral nervous system; PRV, porcine pseudorabies virus; PSA, polysialic acid; SH2, Src homology 2; SH3, Src homology 3; SRCR, scavenger receptor cysteine‐rich; SVZ, subventricular zone; TAM, tyrosine‐based activation motif; VASE, variable alternative‐spliced exon; VSIG, V‐set and immunoglobulin domain containing; WGA, wheat germ agglutinin Protein Names: Abl, v‐abl Abelson murine leukemia viral oncogene homolog; ADAM, a disintegrin and metalloproteinase; ADIP, afadin DIL domain‐interacting protein; ALCAM, activated leukocyte‐cell adhesion molecule; Alx4, Aristaless‐like homeobox 4; AMICA, adhesion molecule, interacts with CXADR antigen 1; AMPA‐R, a‐amino‐3‐hydroxy‐5‐methyl‐4‐isoxazole propionic acid receptor; AP‐2, transcription factor AP‐2 a (activating enhancer‐binding protein 2a); Barx2, BarH‐like homeobox 2; BDNF, brain‐derived neurotrophic factor; Brn‐2, brain‐2 (POU domain, class 3, transcription factor 2); C1‐TEN, tensin‐like C1 domain containing phosphatase, TENC1; CAM, cell adhesion molecule; CaMKII, calcium/calmodulin‐dependent protein kinase II; cAMP, cyclic adenosine monophosphate; CASK, calcium/calmodulin‐associated serine/threonine kinase; Caspr, contactin‐associated protein 1, CNTNAP1; CBP, CREB‐binding protein; CHD, CAM homology domain; CHL1, cell adhesion molecule with homology to L1CAM (close homolog of L1); CKII, casein kinase II; CLMP, coxsackie‐ and adenovirus receptor‐like membrane protein; CEPU‐1, from cerebellar Purkinje cells; CNR, cadherin‐related neuronal receptors; CREB, cyclic‐AMP response‐element binding protein; CSF‐1R, colony‐stimulating factor 1 receptor; Csk, c‐src tyrosine kinase; CTX, cortical thymocyte marker of Xenopus; Cux, cut‐like 1, CCAAT displacement protein, CUTL1; CXADR, coxsackie virus and adenovirus receptor; DAL‐1, differentially expressed adenocarcinoma of the lung, 4.1B, erythrocyte membrane protein band 4.1‐like 3, EPB41L3; DCC, deleted in colorectal cancer; DNAM‐1, DNAX accessory molecule‐1, CD226 antigen; Dock, Dreadlocks (Drosophila); DSCAM, Down syndrome cell adhesion molecule; DSCAML1, Down syndrome cell adhesion molecule‐like 1;

41

42

2


EGF, epidermal growth factor; EGFR, epidermal growth factor receptor; ERK, extracellular signal‐regulated kinase; ESAM, endothelial cell adhesion molecule; Ets‐1, v‐ets erythroblastosis virus E26 oncogene homolog 1; FAK, focal adhesion kinase; FGF, fibroblast growth factor; FGFR, fibroblast growth factor receptor; FLRT1‐3, fibronectin leucine‐rich transmembrane protein 1–3; Fos, v‐fos FBJ murine osteosarcoma viral oncogene homolog; Fps, feline sarcoma oncogene, FES; Fyn, Fyn oncogene related to Src, Fgr, Yes; GAP‐43, growth‐associated protein 43; GDNF, glial cell line‐derived neurotrophic factor; GFAP, glial fibrillary acidic protein; GFRa, GDNF family receptor a; GH, growth hormone; Gli1, glioma‐associated oncogene homolog 1 (zinc finger protein); Gli3, GLI‐Kruppel family member GLI3 (Greig cephalopolysyndactyly syndrome); GRA33, glycoprotein A33; Grb2, growth factor receptor‐bound protein 2; GRIP, glutamate receptor‐interacting protein 1; GSK3b, glycogen synthase kinase‐3b; HGF/SF, hepatocyte growth factor/ scatter factor; Hoxb8, homeo box B8; Hoxb9, homeo box B9; Hoxc6, homeo box C6; IAP, integrin‐ associated protein; ICAT, inhibitor of b‐catenin and Tcf‐4; IGSF9, immunoglobulin superfamily member 9; IgLON, immunoglobulin superfamily containing LAMP, OBCAM, and neurotrimin, IgSF, immunoglobulin superfamily; Inl, internalin; IQGAP, IQ motif‐containing GTPase‐activating protein; IR, insulin receptor; JAK2, Janus kinase 2; JAM, junctional adhesion molecules; JNK, c‐Jun N‐terminal kinase; Kilon, kindred of IgLON; L1, L1 cell adhesion molecule, L1CAM; LAMP, limbic system‐associated membrane protein; LANP, acidic leucine‐rich nuclear phosphoprotein 32 family member A; Lck, lymphocyte‐specific protein tyrosine kinase; LEF‐1, lymphoid enhancer‐binding factor 1; LMO7, LIM domain 7; LNX, ligand‐of‐Numb protein‐X; Lyn, v‐yes‐1 Yamaguchi sarcoma viral‐related oncogene homolog; MAG, myelin‐associated glycoprotein; MAGI‐1b, membrane‐associated guanylate kinase interacting protein‐ like 1; MAP, mitogen‐activated protein; MAP1A, microtubule‐associated protein 1A; MAP2, microtubule‐ associated protein 2; MAPK, mitogen‐activated protein kinase; MCAM, melanoma cell adhesion molecule; MEK, mitogen‐activated protein kinase kinase; MERTK, c‐Mer protooncogene tyrosine kinase; MFR, macrophage fusion receptor; MITF, microphthalmia‐associated transcription factor; mLin‐7, lin‐7 homolog A, LIN7A; MMP, matrix metalloproteinases; MPDZ, multiple PDZ domain protein; MSK1, mitogen‐ and stress‐activated protein kinase 1; NCAM, neural cell adhesion molecule; Nck, NCK adaptor protein 1; Necl, nectin‐like protein; NF‐kB, nuclear factor‐kappa B; NgCAM, neuron–glia cell adhesion molecule; NGF, nerve growth factor; Nogo‐66; reticulon 4 receptor, RTN4R; Nope, neighbor of Punc E11; NrCAM, NgCAM‐related cell adhesion molecule, neuronal cell adhesion molecule; NT‐3, neurotrophin‐3; OBCAM, opioid‐binding cell adhesion molecule; OCAM, olfactory cell adhesion molecule; Otx2, orthodenticle homolog 2; PACSIN1, protein kinase C and casein kinase substrate in neurons protein 1; Pak, p21/Cdc42/Rac1‐activated kinase 1 (STE20 homolog, yeast); Pals2, membrane protein, palmitoylated 6 (MAGUK p55 subfamily member 6); PAR‐3, partitioning‐defective 3 homolog; PARC, paraxial protocadherin; Pax2, paired box gene 2; Pax3, paired box gene 3 (Waardenburg syndrome 1); Pax6, paired box gene 6 (aniridia, keratitis); Pax8, paired box gene 8; Phox2, paired‐like homeobox 2; PCR, polymerase chair reaction; PDGF, platelet‐derived growth factor; PDGFR, platelet‐derived growth factor receptor; PDK, phosphoinositide‐dependent kinase; PDZ, postsynaptic density‐95 (PSD‐95)/discs large/zona occludens‐1; PI3K, phosphatidylinositol‐3‐kinase; PICK1, protein interacting with C‐kinase‐1 (PKC, a binding protein, PRKCABP); PKA, protein kinase A; PKB, protein kinase B; PKC, protein kinase C; PLC, phospholipase C; PLD, phospholipase D; PMP22, peripheral myelin protein 22; PP1, protein phosphatase 1, PP2A, protein phosphatase 2A; PS1, presenilin 1; PSD‐95, postsynaptic density‐95 (discs, large homolog 4, DLG4); PTP, protein tyrosine phosphatase; PTPNS1, protein tyrosine phosphatase, nonreceptor type substrate 1; PTPNS1L, protein tyrosine phosphatase, nonreceptor type substrate 1‐like; Punc, putative neuronal cell adhesion molecule; PVR, poliovirus receptor; PYK2, PTK2B protein tyrosine kinase 2b; Rac, Ras‐related C3 botulinum toxin substrate; Rag‐1, recombination‐activating gene 1; RAGE, renal tumor antigen; RanBPM, Ran‐binding protein 9; rhBMP‐2, recombinant human bone morphogenetic protein‐2; Rho, Ras homolog gene family member; RNCAM, Rb‐8‐neural cell adhesion molecule; Robo1–4; roundabout, axon guidance receptor, homolog 1–4 (Drosophila); ROKa, Rho A‐binding kinase a; RPTP, receptor‐like protein tyrosine phosphatase; Rsk, ribosomal protein S6 kinase; SAP97, discs, large homolog 1 (Drosophila), DLG1; SAP102, synapse‐associated protein 102; Sdk1‐2, sidekick homolog 1–2 (chicken); Shank, Src homology 3 domain and ankyrin repeat‐containing protein; Shc, Src homology 2 domain‐containing transforming


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protein 1; Shh, Sonic hedgehog; SHP, SH2‐containing tyrosine phosphatase; SHP‐2, protein tyrosine phosphatase, nonreceptor type 11, PTPN11; SHPS‐1, SHP substrate 1; SIRP, signal‐regulatory protein; SKAP55hom/R, Src‐kinase‐associated protein of 55‐kDa homologue; Slit1–3; Slit homolog 1–3 (Drosophila); Socs‐1, suppressor of cytokine signaling 1; Src, v‐src sarcoma (Schmidt‐Ruppin A‐2) viral oncogene homolog; SrGAP, Slit–Robo Rho GTPase‐activating protein; S‐SCAM, synaptic scaffolding molecule; Tctex‐1, t‐complex testis‐expressed gene 1; TMEM25, transmembrane protein 25; TOAD‐64, turned on after division‐64; Vav1, vav 1 oncogene; VLA‐5, integrin a5b1; Wnt, Wingless; ZO‐1, zona occludens 1 (tight junction protein 1, TJP1)

1

Introduction

This chapter deals with cell adhesion molecules (CAMs) of the nervous system belonging to the immunoglobulin superfamily, IgSF. The majority of the presented CAMs are not exclusively expressed in the nervous system, but the descriptions given in this chapter focus specifically on the expression and function of the CAMs in the nervous system. Hence, some of the molecules may in other tissues have important functions, which are not mentioned, and the individual presentations—and the literature cited—are therefore not always representative of the overall function of the described proteins in the organism in general.

1.1 Defining an Adhesion Molecule Adhesion molecules have traditionally been divided into four groups: the cadherins, the selectins, the integrins, and molecules of the IgSF. According to some authors, the phrase ‘‘cell adhesion molecule’’ includes all proteins mediating adhesion, whether it is between cells (cell–cell interactions) or between cells and the extracellular matrix (ECM) (cell–ECM interactions). Other authors restrict CAMs to include proteins mediating cell–cell interactions, whereas the remaining adhesion molecules are referred to as ‘‘substrate adhesion molecules.’’ The more restricted definition of CAMs is understandable, but leaves the problem that many molecules can be classified as both CAMs and substrate adhesion molecules. In addition, not a single adhesion molecule seems to be expressed with the sole purpose of mediating adhesion. Furthermore, cellular adhesion is not an isolated phenomenon. It has been demonstrated that adhesion in itself is sufficient to modulate gene transcription, and consequently, the expression of adhesion molecules typically affect intracellular signaling, and modulate cell survival, proliferation, and differentiation (Ingber, 2003). Under some circumstances adhesion molecules even serve to reduce intercellular adhesion (as for instance NCAM expressing the rare type of glycosylation known as polysialylation) (see later). Thus, the mediation of adhesion is one of several functions of an adhesion molecule, and it may not always be the most significant function of the protein.

1.2 Restrictions in Protein Types and Species The chapter predominantly describes the function of mammalian proteins, and the tables accompanying the individual sections only include genetic information for the proteins in humans and mice. In these tables, the ‘‘HGNC ID’’ numbers refer to the database IDs of the ‘‘HUGO Gene Nomenclature Committee’’ (http://www.gene.ucl.ac.uk/nomenclature/), whereas ‘‘MGI ID’’ numbers refer to the database IDs at the ‘‘Mouse Genome Informatics’’ (http://www.informatics.jax.org/). Finally, ‘‘Locus ID’’ numbers refer to the database IDs at ‘‘Locus Link’’ at the National Center for Biotechnology Information (NCBI; http://www. ncbi.nlm.nih.gov/LocusLink/). These sites all contain numerous links to other pages with additional information of the indicated genes and proteins. Most proteins are known by several names. Hence, each CAM description begins with a list of synonyms including abandoned gene symbols. No typographical distinctions have been made between proteins from

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different mammalian species. However, in most situations it will be apparent from the text, whether a given study has been performed with the mouse, rat, or human orthologue of a given protein. In this chapter the selection of proteins has been restricted only to include molecules that mediate cell– cell interactions. Some of the described proteins are ‘‘canonical’’ CAMs, whereas the ability to mediate adhesion for some of the other presented proteins only seems to be a minor part of their overall function. At the end of the chapter we also present a few ‘‘putative’’ adhesion molecules, for which the ability to mediate intercellular adhesion can be assumed although it has not been demonstrated so far. Importantly, the number of CAMs in the nervous system is large, and in addition, many new proteins have been cloned within recent years. Therefore, it is not the intention here to include all recognized or putative adhesion molecules of the nervous system, but to give an impression of the variability in the structure and function of CAMs in the nervous system. Integrins predominantly mediate cell–substrate interactions and are therefore not included. (For recent reviews of integrin function in the nervous system see Gall and Lynch (2004), Nakamoto et al. (2004), and Clegg et al. (2003)). Selectins mediate binding of activated endothelial cells to leukocytes. Thus, they mainly exert their function outside the nervous system and are therefore also excluded here.

1.3 Cis‐Interactions, Trans‐Interactions, and Homophilic and Heterophilic Interactions CAMs are often able to form dimers, tetramers, or polymers, where one molecule of a given CAM interacts with one or several other copies of the same protein. Such interactions are referred to as homophilic interactions. In addition, most CAMs interact with other molecules—heterophilic interactions—with either their extracellular and/or intracellular domains. CAMs are involved in homo‐ or heterophilic interactions with molecules positioned on opposing plasma membranes, and such interactions are referred to as trans‐interactions. In addition, CAMs are often involved in homo‐ or heterophilic interactions with other membrane‐associated molecules positioned in the same plasma membrane. Such membrane‐lateral interactions are referred to as cis‐interactions. In this chapter, the descriptions of the individual CAMs are supplemented with tables of homophilic and heterophilic binding partners, and often it is indicated whether a given interaction is a cis‐ and/or a trans‐interaction.

1.4 The Immunoglobulin Superfamily The Ig‐like CAMs constitute a subgroup of the IgSF, which is one of the largest families of proteins in vertebrates (in humans, the IgSF has around 765 members) (see Bru¨mmenforf and Lemmon (2001)). Ig‐CAMs contain at least one extracellular module (EC module) with homology to the modules of immunoglobulins. In addition, Ig‐CAMs often contain one or more of the closely related fibronectin type III (F3)‐homology modules, Fn3, and they may also contain other EC modules. Ig‐homology modules (Ig‐modules) are 70‐ to 110‐amino acid‐long globular structures formed by 7–9 antiparallel b‐strands arranged in a sandwich of two b‐sheets with 3–5 b‐strands in each sheet. Furthermore, the modules contain a highly conserved disulfide bridge (some modules contain more than one). The modules are often encoded by two exons and can be divided into four different types, the C1‐ and C2‐types (C for constant), the V‐type (V for variable), and the I‐type (I for intermediate). The V‐type Ig‐modules contain nine b‐strands and have 65–75 amino acids between the conserved disulfide bridges, whereas the C‐types only contain seven b‐strands and have only 55–60 amino acids between the disulfide bridge (Williams and Barclay, 1988). The C1‐type modules are almost exclusively found in immunoglobulins and MHC antigens. The C2‐type modules are structurally related to the C1‐type, but share a stronger sequence homology to the V‐types (Barclay, 2003). The I‐type module is an intermediate between the C‐ and V‐type modules, and it is structurally closest to the V‐type (Harpaz and Chothia, 1994). The Ig‐modules were originally identified in immunoglobulins, and hence the name. However, evolutionarily the CAMs of the IgSF are predecessors of the immunoglobulins of the immune system. Ig‐modules


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do not possess any enzymatic activities, but the modules are capable of establishing relatively weak interactions with other molecules (or each other). The loops combining the individual b‐strands have the potential for a large sequence variability, which can be used to modulate the interactions. The Fn3‐modules are—together with cadherin modules and cytokine receptor modules—structurally closely related to Ig‐modules (Barclay, 2003). The Fn3‐modules most closely resemble Ig‐modules of the C2‐type, containing the same number of b‐strands (seven) organized in a sandwich of two b‐sheets. However, Fn3‐modules do not contain the strongly conserved cysteine residues found in Ig‐modules, and thus rarely contain disulfide bridges (Bru¨mmendorf and Rathjen, 1996). Proteins containing Fn3‐modules also constitute a large superfamily, but when two Fn3 modules are compared, the level of sequence identity within the family can be as low as 5%. However, the module contains a number of conserved residues including some prolines, which have been shown to be important for the modulation of intermodule interactions (Steward et al., 2002). Ig‐CAMs can be classified in several ways. They are often—on the basis of sequence homologies and functional similarities—placed in subfamilies, and individual members of subfamilies are described together here. Another way of categorizing Ig‐CAMs is to group them according to the number of Ig‐ and Fn3‐modules and the sequence in which they are organized. Thus, nectins, necls, and members of the IgLON family are all members of the 3/0 subfamily, whereas F3 and TAG‐1 are members of the 5/4 subfamily (see > Figure 2-1) (Cunningham, 1995). In this chapter, the Ig‐CAMs are presented in the order of increasing structural complexity as indicated in > Figure 2-1, starting with molecules containing a few Ig‐modules and finishing with molecules containing multiple Ig‐ and Fn3‐modules. However, a few of the molecules shown in > Figure 2-1 are described in a separate section as ‘‘putative adhesion molecules,’’ since their involvement in cell–cell interactions is unknown. For additional reviews of IgSF CAMs, see Bru¨mmendorf and Rathjen (1996), Bru¨mmendorf and Lemmon (2001), Chothia and Jones (1997), and Rougon and Hobert (2003).

2

Myelin Protein Zero

2.1 Introduction Myelin protein zero (P0) was described as a major component of peripheral myelin in the 1970s (Brostoff et al., 1975). However, the genes encoding rodent P0 were not cloned until the 1980s (Lemke and Axel, 1985; Lemke et al., 1988). In 1991, full‐length P0 protein was isolated from human spinal cord and the deduced amino acid sequence was found to be very homologous to the rodent sequences (Hayasaka et al., 1991), encoding a transmembrane protein with a single extracellular Ig‐module. The amino acid sequence of the extracellular domain of P0 is 46% identical (60% similar) to the more recently cloned myelin protein zero‐like 1 (MPZL1). However, MPZL1is only expressed in low amounts in the nervous system (Zhao and Zhao, 1998) and will not be described further here. See also > Table 2-1.

2.2 Genes The approved name of the gene encoding P0 is MPZ (myelin protein zero) (> Table 2-2). The gene contains six exons and spans 6–7 kb (Lemke et al., 1988; You et al., 1991; Hayasaka et al., 1993; Pham‐Dinh et al., 1993). The protein exists in several isoforms as a result of alternative splicing of the gene product (Besançon et al., 1999). The expression of P0 has been demonstrated to be induced by the transcription factor Sox10 (Peirano et al., 2000).

2.3 Expression The expression of P0 is restricted to Schwann cells in the peripheral nervous system (PNS). The expression of P0 roughly parallels the process of myelination and P0 accounts for more than 50% of the protein present in the sheath of peripheral nerves (Lemke, 1986).

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. Figure 2-1 Cell adhesion molecules (CAMs) of the immunoglobulin superfamily (IgSF). The figure shows the modular organization of the CAMs described in the text. The approximate length of the cytoplasmic tails of transmembrane proteins can be read from the scalebar at the bottom of the figure. I, V, C1, and C2 indicate the Ig‐module type


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. Table 2-1 Nomenclature for P0 and PZR CAM name P0 PZR

Protein and gene synonyms MPP, myelin peripheral protein, P‐zero, P>0zero< Myelin protein zero‐like 1, PZR, FLJ21047

. Table 2-2 Gene information for P0 and PZR

Species Man

Mouse

Approved gene symbol MPZ MPZL1 Mpz Mpzl1

Approved gene name Myelin protein zero (Charcot–Marie–Tooth neuropathy 1B) Myelin protein zero‐like 1 Myelin protein zero Myelin protein zero‐like 1

Locus 1q22–q23

Database reference HGNC ID: 7225

1q24.2 1 92.4 1 syntenic

HGNC ID: 7226 MGI ID: 103177 MGI ID: 1915731

During myelination of neural cells, NCAM and L1 are present on neurons and Schwann cells prior to myelination, but downregulated as myelination starts. This is roughly the time point where P0 and MAG become detectable (Martini et al., 1995). Before myelin compaction, the subcellular location of P0 and MAG is identical. However, after compaction MAG and P0 are located in a complementary manner, where MAG is confined to noncompacted areas of the Schwann cells and P0 is restricted to the compacted areas (see also > Sect. 13) (reviewed by Martini et al. (1994)).

2.4 Isoforms and Protein Structure P0 is structurally the simplest member of the IgSF. It contains a single extracellular Ig‐module of the V‐type followed by a transmembrane segment and an 69‐amino acids‐long highly charged cytoplasmic tail. Given the simple protein structure, P0 has been suggested to be a close relative to the ancestral gene from which the IgSF has developed (You et al., 1991). Alternative splicing of MPZ can result in truncated isoforms of P0. These truncated isoforms are the result of partial or complete removal of exon three and a simultaneous truncation of the product owing to a frameshift that leads to a premature stop codon in exon four (Besançon et al., 1999).

2.5 Posttranslational Modifications P0 contains a single site for N‐glycosylation, and this site is occupied by a carbohydrate with a HNK‐1 epitope (an isotope expressed on a number of CAMs, including NCAM, L1, TAG‐1, and PTPNS1) (Kleene and Schachner, 2004). P0 may also be acetylated at C153, and phosphorylated at the cytoplasmic residues S191 and S204, and it also contains a consensus sequence for tyrosine phosphorylation (Eichberg, 2002).

2.6 Homophilic Interactions The structure of the extracellular Ig‐module of P0 has been determined by X‐ray crystallography. The structure indicates that P0 forms tetramers by cis‐interactions. These tetramers may then mediate trans‐ interactions with tetramers on opposing membranes. Alternatively, Trp residues extending from the tetramers may mediate trans‐cellular interactions by being buried within the lipid bilayer of opposing membranes (> Figure 2-2; > Table 2-3) (Shapiro et al., 1996; Martini and Carenini, 1998).

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. Figure 2-2 Myelin Protein Zero (P0) (a): Ribbon diagram of the X‐ray structure of the Ig‐module of P0. The module is of the V‐type. (b, c): Putative organization of P0 molecules in the extracellular space between two cell membranes. The figure shows five tetramers of P0; four (yellow) protruding from the lower membrane in (b), and one (blue) emanating from the upper membrane in (b). The highlighted regions in (b) show the linker regions connecting the respective Ig‐modules to the plasma membrane, and the Trp‐residues that may link the Ig‐modules to the opposing plasma membrane (Taken from Shapiro et al., 1996)

. Table 2-3 Binding partners for P0 CAM P0 P0

Extracellular binding Partner P0 (cis, trans) Plasma membrane phospholipids peripheral myelin protein 22

Reference Shapiro et al. (1996); Martini and Carenini, (1998) Martini and Carenini (1998) D’Urso et al. (1999)


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2.7 Heterophilic Binding Partners The cytoplasmic tail of P0 has been suggested to interact with phospholipids in the plasma membrane (reviewed by Martini and Carenini (1998)). Immunoprecipitation studies suggest cis and/or trans‐interactions between P0 and another transmembrane receptor, the peripheral myelin protein 22 (PMP22) (D’Urso et al., 1999) (> Table 2-3).

2.8 Function P0 promotes myelination and plays a pivotal role in both myelin formation and maintenance. This has been demonstrated in P0‐knockout mice, which display severe hypomyelination in the PNS (reviewed by Martini and Carenini (1998)), and P0 knockouts also demonstrate dysregulation of the expression of voltage‐gated sodium channels in the nodes of Ranvier (Ulzheimer et al., 2004). In humans, P0 mutations are associated with peripheral neuropathies, among which are Charcot– Marie–Tooth disease, De´je´rine–Sottas syndrome, and congenital hypomyelination (for review, see Eichberg (2002)).

3

Integrin‐Associated Protein

3.1 Introduction Integrin‐associated protein (CD47) was originally identified and isolated by several research groups and has consequently been given several names including integrin‐associated protein (IAP) (Brown et al., 1990), OA3 antigen (Campbell et al., 1992), and CD47 (Mawby et al., 1994) (> Table 2-4). . Table 2-4 Nomenclature for CD47 CAM name CD47

Protein and gene synonyms MER6, IAP, integrin‐associated protein, 9130415E20Rik, B430305P08Rik, Itgp, antigenic surface determinant protein OA3

The protein has an unusual structure; a single Ig‐module followed by a so‐called multiple membrane‐ spanning segment with five transmembrane regions. It is widely expressed by hematopoietic and lymphoid cells (Mawby et al., 1994), where it has been related to the modulation of cytokine synthesis. Furthermore, extracellular interactions with CD47 can induce cell death through the stimulation of the heterotrimeric G protein Gia and subsequent inhibition of protein kinase A (see Pettersen, 2000; Brown and Frazier, 2001; Manna and Frazier, 2003; for reviews). However, CD47 is also expressed in neuronal tissues, where it is involved in neurite outgrowth, synaptogenesis, and memory formation (Lee et al., 2000a; Numakawa et al., 2004; Ohnishi et al., 2005).

3.2 Gene In humans, CD47 has been mapped to chromosome 3, where the gene spans 45 kb (Schickel et al., 2002) (> Table 2-5). The human CD47 promoter contains a binding site for the transcription factor a‐Pas/NRF‐1 (a transcription factor regulating proteins involved in e.g. cellular proliferation), and overexpression of this factor enhances the expression of CD47 (Chang and Huang, 2004).

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. Table 2-5 Gene information for CD47 Species Man

Approved gene symbol CD47

Mouse

Cd47

Approved gene name CD47 antigen (Rh‐related antigen, integrin‐associated signal transducer) CD47 antigen (Rh‐related antigen, integrin‐associated signal transducer)

Locus 3q13.1–q13.2


16 syntenic

MGI ID: 96617

Human CD47 contains 12 exons (named 1 to 7, EN (novel exon), Form2, Form3, Form4, and 30 ‐UT). Alternative splicing of CD47 results in at least five different isoforms (Reinhold et al., 1995; Schickel et al., 2002).

3.3 Expression CD47 is widely expressed in lymphoid and hemopoietic cells. However, the protein is also expressed in several other tissues including placenta, liver, pancreas, and brain (Brown et al., 1990; Mawby et al., 1994). Neural tissues predominantly express the so‐called isoform 4 (see below), which also is expressed in liver and testis (Reinhold et al., 1995). However, in rat, isoforms 2 and 3 have also been detected in neurons (Huang et al., 1998). In cultured hippocampal neurons, CD47 has been demonstrated predominantly to be localized at the surface of dendrites (Ohnishi et al., 2005). In the mouse retina, CD47 and its counter‐receptor, PTPNS1, localize to synapses. Interestingly, the expression of CD47 precedes the expression of PTPNS1, and the synaptic localization of PTPNS1 is abrogated in CD47‐knockout mice (Mi et al., 2000).

3.4 Isoforms and Protein Structure CD47 is 323 amino acids long and contains a single Ig‐module of the V‐type followed by a multiple membrane‐spanning segment containing five transmembrane regions and a short (up to 33 amino acids long) cytoplasmic tail (Campbell et al., 1992). Five different isoforms of the protein have been identified (named 1, 2, 3, 4, and new) of which one (new) is reported to be specifically expressed in muscle. All identified isoforms contain the Ig‐module and the multiple membrane‐spanning segment, but differ in the length of the cytoplasmic tail (isoform 1 having the shortest and isoform 4 the longest tail) (> Figure 2-3) (Reinhold et al., 1995; Schickel et al., 2002).

3.5 Posttranslational Modifications Human CD47 contains six potential sites for N‐linked glycosylations (Mawby et al., 1994). Furthermore, the protein contains an unusual disulfide bridge linking the Ig‐module with the extracellular loop located between the fourth and fifth transmembrane segments. Abrogation of this disulfide bridge reduces the extracellular protein interactions mediated by CD47, and affects signaling mediated by the protein (Rebres et al., 2001).

3.6 Homophilic Interactions CD47 has not been reported to form homophilic interactions.


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. Figure 2-3 Integrin‐Associated Protein (CD47). General model of the structure of CD47. The protein consists of a single Ig‐module (blue) followed by five transmembrane regions (orange) with interconnecting loops (red) and a cytoplasmic tail (green). Alternative splicing results in CD47 isoforms with four different lengths of the cytoplasmic tail (indicated by 1–4). The Ig‐module is crosslinked to the membrane spanning domain by a disulfide bridge (S–S) (Taken from Oldenborg, 2004)

3.7 Heterophilic Binding Partners 3.7.1 Extracellular Binding Partners CD47 forms cis‐interactions with several types of integrins (Brown et al., 1990), including the fibrinogen receptor aIIbb3 (Fujimoto et al., 2003), the collagen receptor a2b1 (Wang and Frazier, 1998), the RGD‐ binding avb3 (McDonald et al., 2004), and the fibronectin receptor a4b1 (Barazi et al., 2002). Moreover, CD47 cis‐interacts with the surface receptor Fas/CD95, an interaction important for Fas‐mediated apoptosis (Manna et al., 2005). CD47 also interacts with several proteins of the ECM. Thus, it binds the C‐terminal domain of thrombospondin‐1 (Gao et al., 1996), a large secreted protein, which interacts with a number of ECM proteins and is involved in several biological processes including angiogenesis and platelet aggregation. There are five types of thrombospondin (also known as TSP‐1 to ‐5), and CD47 is believed to bind all five (see Brown and Frazier, 2001). Furthermore, it binds the a3 chain of collagen IV in the basement membrane, an interaction reported to affect the proliferation of melanoma cells (Shahan et al., 1999). CD47 has been shown to form trans‐interactions with the two SIRP family receptors SIRPB2 (Brooke et al., 2004; Piccio et al., 2005) and PTPNS1 (Jiang et al., 1999; Seiffert et al., 1999; Vernon‐Wilson et al., 2000) (> Table 2-6). See Sect. 8 on the SIRP family for a more detailed description of the CD47–PTPNS1

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. Table 2-6 Binding partners for CD47 CAM CD47

Extracellular binding partner PTPNS1 (trans)

CD47

SIRPB2 (trans) Collagen IV Thrombospondin‐1 aIIbb3 (cis) avb3(cis) a2b1 (cis) a4b1 (cis) Fas/CD95 (cis) Ubiquilin 1, ubiquilin 2

Reference Jiang et al. (1999); Seiffert et al. (1999); Vernon‐Wilson et al. (2000); Han et al. (2000) Piccio et al. (2005) Shahan et al. (1999) Gao et al. (1996) Fujimoto et al. (2003) McDonald et al. (2004) Wang and Frazier (1998) Barazi et al. (2002) Manna et al. (2005) Wu et al. (1999)

interaction. Moreover, CD47 has been reported to modulate cellular migration and intercellular adhesion through the trans‐interaction with unidentified counter receptor(s) (Rebres et al., 2005).

3.7.2 Intracellular Binding Partners The cytoplasmic tail of CD47 binds ubiquitin‐like proteins ubiquilin 1 (also known as PLIC‐1) and ubiquilin 2 (also known as CHAP1 and PLIC2) (Wu et al., 1999) (> Table 2-6). These proteins bind to the proteasome and to ubiquitin ligases (E3 enzymes) and have been shown to interfere with proteasome‐ mediated degradation of the tumor suppressor protein p53 (Kleijnen et al., 2000). Furthermore, they may link CD47 to intermediate filaments (Wu et al., 1999). In human erythrocytes, CD47 has been suggested to link the band 3 cytoskeletal complex with Rhesus protein complexes (see Oldenborg, 2004).

3.8 Function CD47 is important for learning and memory. Thus, in vivo experiments have demonstrated that CD47 is expressed in higher amounts in rats with good memory compared with rats with poor memory, and three hours subsequent to a memory test (a one‐way inhibitory passive avoidance learning task) rats with good memory demonstrated strong increase in CD47 expression. Furthermore, the expression of the protein is increased in response to injection of NMDA (N‐methyl‐D‐aspartate) or amphetamine into the hippocampus (Huang et al., 1998). Isoforms of CD47 increased during learning and memory consolidation include isoforms 3 and 4—but not isoform 2 (Lee et al., 2000a). Moreover, knockdown of CD47 in rats and knockout of CD47 in mice leads to a significant impairment in memory consolidation (Huang et al., 1998; Chang et al., 1999). In vitro, overexpression of CD47 (isoform 4) in cultured cortical neurons leads to significant increase in neurite outgrowth and subsequent increase in glutamate‐mediated neurotransmission and upregulation of synaptic markers, including synaptotagmin, syntaxin, synapsin I, and SNAP25. The effects are dependent on CD47‐mediated activation of the MAPKs, ERK1 and ‐2, whereas the PI3K–PKB pathway and the activity of the TrkB receptor are unaffected (Numakawa et al., 2004). Surprisingly, overexpression of CD47 in cortical neurons has also been reported to induce cell death by apoptosis, an effect enhanced by coexpression of the counter‐receptor, PTPNS1, but reduced by treatment with the growth factor BDNF (Koshimizu et al., 2002).


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In neuroblastoma cells, CD47 can stimulate neurite outgrowth and filopodia formation through different signal transduction pathways. Thus, filopodia formation involves the activation of the small GTPase Cdc42 and requires the presence of intact CD47, whereas induction of neurite outgrowth involves the activation of the small GTPase Rac and requires the Ig‐module of CD47 and the interaction between CD47 and integrin b3 (Miyashita et al., 2004). In addition to its effects in cells of neuronal origin, CD47 is involved in macrophage fusion (Saginario et al., 1998; Han et al., 2000), intercellular interactions mediated by T cells (Latour et al., 2001; Piccio et al., 2005), chemotaxis and transmigration of neutrophils (Liu et al., 2002), and transmigration of monocytes across the cerebral endothelium, a process of central importance for the development of neuroinflammatory diseases (de Vries et al., 2002). For reviews of CD47 see Brown and Frazier (2001), Oldenborg (2004), and Pettersen (2000).

4

The CTX Family

4.1 Introduction The CTX family (CXADR, IGSF11) is constituted by 13 proteins and is named after a molecule, CTX, which is highly expressed in cortical thymocytes in Xenopus. The mouse and human homologs of CTX were cloned in 1998 (Chre´tien et al., 1998). They were originally named CTM and CTH, respectively, but the official gene symbols are Vsig2/VSIG2. The CTX family also includes the proteins AMICA, ASAM, ESAM, CXADR, GRA33, IGSF11, VSIG1, VSIG2, and VSIG4 and the subfamily of JAM proteins (F11R, JAM‐2, JAM‐3, and JAM‐4) (> Table 2-7). . Table 2-7 Nomenclature for CTX Family CAM name AMICA ASAM CXADR ESAM F11R GRA33 IGSF11 JAM2 JAM3 JAM4 VSIG1 VSIG2 VSIG4

Protein and gene synonyms Adhesion molecule, interacts with CXADR antigen 1, Gm638, JAML, junctional adhesion molecule‐like, LOC270152 ACAM, CLMP, CAR‐like membrane protein, RIKEN cDNA 9030425E11 gene 2610206D03Rik, CAR 2310008D05Rik, W117m PAM‐1, JCAM, JAM‐1, junctional adhesion molecule‐1, JAM‐A, BV11 antigen, ESTM33, F11R, JAM, Jcam1, Ly106 A33, 2010310L10Rik, 2210401D16Rik BT‐IgSF, MGC35227, VSIG3 VE‐JAM, HGNC:1284, JAM‐B, 2410030G21Rik, 2410167M24Rik, JAM‐2, Jcam2 JAM‐C, 1110002N23Rik, JAM‐3, Jcam3 2010003D20Rik, Igsf5, JCAM 1700062D20Rik, MGC44287 1190004B15Rik, 2210413P10Rik, CTH, CTM, CTX, CTXL Z39IG

The majority of the proteins constituting the CTX family is expressed in limited amounts in the nervous system and are only described briefly. AMICA was cloned in 2003 and is mainly expressed in hematopoietic tissues (Moog‐Lutz et al., 2003). ASAM (CLMP), the most recently cloned member of the CTX family, is expressed by adipocytes (Eguchi et al., 2005) and epithelial cells, where it colocalizes with the tight junction proteins ZO‐1 and occludin in cell–cell contacts and induces cellular aggregation. In humans, it is mainly found in the intestine and placenta, whereas in the mouse it is most strongly expressed in the brain and

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heart (Raschperger et al., 2004). ESAM was cloned in 2001 by Hirata and coworkers. It is involved in angiogenic processes and is expressed in vascular endothelial cells, predominantly in the lung and heart (Hirata et al., 2001; Ishida et al., 2003). GRA33 is expressed in >95% of human colon cancers but is absent from most other human tissues and tumor types (Heath et al., 1997). The expression of VSIG1 has not been investigated. VSIG2 n humans is found in various tissues, including stomach, colon, and prostate. In the mouse, the protein is predominantly found in prostate and stomach (Chre´tien et al., 1998). VSIG4 in humans is found in most tissues, but predominantly in lung and placenta, and in synovium from patients with rheumatoid arthritis (Langnaese et al., 2000; Walker, 2002). The protein has also been found in subsets of dendritic cells (Ahn et al., 2002). The JAM family consists of F11R, JAM2, JAM3, and JAM4 (Martin‐Padura et al., 1998; Ozaki et al., 1999; Arrate et al., 2001; Aurrand‐Lions et al., 2001; Naik et al., 2001; Liang et al., 2002; Hirabayashi et al., 2003; Moog‐Lutz et al., 2003). This group of proteins is expressed by endothelial cells, epithelial cells, leukocytes, and platelets, where they are involved in tight junction formation, endothelial migration, and transendothelial migration of leukocytes (Martin‐Parura et al., 1998; Johnson‐Le´ger et al., 2002; Liang et al., 2002; Naik et al., 2003). The nomenclature for JAMs has been proposed to be changed from JAM1‐4 to JAMA‐D (Bazzoni, 2003).

4.2 CXADR 4.2.1 Gene Locus information for CXADR is given in > Table 2-8. Human and mouse CXADR were cloned in 1997 (Bergelson et al., 1997; Tomko et al., 1997). The human gene has been shown to contain seven exons and the protein exists in five isoforms because of alternative splicing (see below) (Bowles et al., 1999).

4.2.2 Expression CXADR is expressed in several tissues. In humans, the highest expression is found in the brain, heart, pancreas, small intestine, testis, and prostate. In the mouse, the protein is predominantly found in the liver, but also in the brain, heart, kidney, and lung (Tomko et al., 1997). The expression of CXADR is highly temporally regulated. In the mouse, the highest level of CXADR expression is found around birth, after which the amount decreases rapidly, and in the adult mouse brain the protein is absent or expressed in very low amounts. Expression appears in the embryonic ectoderm on E6.5. From E8.5 to postnatal day (P) 7 a strong expression of the protein has been demonstrated in the neuroepithelium of the neural tube, the developing brain, and the spinal cord. From E9.5 to E11.5 it is expressed in the cranial motor nerves, and from E13.5 to P7 it has been detected in the optic nerve. CXADR has also been observed to be expressed in the cell bodies, neurites, and growth cones of cultured E16 hippocampal neurons (Honda et al., 2000; Hotta et al., 2003). Recently, in humans and mice CXADR was shown also to be expressed in the neuromuscular junctions of skeletal muscles and the intercalated discs of cardiac muscles (Shaw et al., 2004). Furthermore, CXADR is expressed in various types of cancers. However, there is a large variation in the degree of CXADR expression between different tumors (Fuxe et al., 2003).

4.2.3 Isoforms and Protein Structure All members of the CTX family have the same general protein structure: two Ig‐modules (one V‐type module and one C2‐type module), a transmembrane segment, and a cytoplasmic tail. The CXADR protein is up to 365‐ and 355 amino acid long in humans and mice, respectively, with a 222‐amino acid‐long extracellular part and a 97‐ to 107‐amino acid‐long cytoplasmic tail


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. Table 2-8 Gene information for CTX family members

Species Man

Approved gene symbol AMICA1 ASAM CXADR ESAM F11R GPA33 IGSF11 JAM2 JAM3 VSIG1 VSIG2 VSIG4

Mouse

Gm638 9030425E11Rik Cxadr Esam1 F11r Gpa33 Igsf11 Jam2 Jam3 Jam4 4930405J24Rik Vsig2

Approved gene name Adhesion molecule, interacts with CXADR antigen 1 Adipocyte‐specific adhesion molecule Coxsackie virus and adenovirus receptor Endothelial cell adhesion molecule F11 receptor Glycoprotein A33 (transmembrane) Immunoglobulin superfamily, member 11 Junctional adhesion molecule 2 Junctional adhesion molecule 3 V‐set and immunoglobulin domain containing 1 V‐set and immunoglobulin domain containing 2 V‐set and immunoglobulin domain containing 4 Ggene model 638, (NCBI) (orthologue to human AMICA1) RIKEN cDNA 9030425E11 gene (orthologue to human ASAM) Coxsackie virus and adenovirus receptor Endothelial cell‐specific adhesion molecule F11 receptor Glycoprotein A33 (transmembrane) Immunoglobulin superfamily, member 11 Junctional adhesion molecule 2 Junctional adhesion molecule 3 Junctional adhesion molecule 4 RIKEN cDNA 4930405J24 gene (orthologue to human VSIG1) V‐set and immunoglobulin domain containing 2

Locus 11q23.3

Database reference HGNC ID: 11q23.3

11q24.1 21q 11q24.2 1q21.2–q21.3 1q23–1q24.1 3q21.2 21q21.2 11q25 Xq22.3

GeneID: 79827 HGNC ID: 2559 HGNC ID: 17474 HGNC ID: 14685 HGNC ID: 4445 HGNC ID: 16669 HGNC ID: 14686 HGNC ID: 15532 HGNC ID: 28675

11q24

HGNC ID: 17149

Xq12–q13.3

HGNC ID: 17032

9 syntenic

MGI: 2685484

9 syntenic

MGI: 1918816

16 C 9B

MGI: 1201679 MGI: 1916774

1 93.3 H2 1 86.2 16 syntenic 16 syntenic 9 syntenic 16 C4 X F1

MGI: 1321398 MGI: 1891703 MGI: 2388477 MGI: 1933820 MGI: 1933825 MGI: 1919308 MGI: 1926039

9B

MGI: 1928009

(Tomko et al., 1997). The human and murine proteins are 91% identical, and the cytoplasmic tails exhibit 95% identity (Bergelson et al., 1997). The two Ig‐modules of CXADR are named D1 and D2, respectively (Bewley et al., 1999). CXADR exists in two transmembrane isoforms of 365‐ and 352 amino acids, and three soluble isoforms of 252‐, 200‐, and 89 amino acids, respectively. The transmembrane isoforms differ only in the length of the cytoplasmic tail and are referred to as SIV and TVV, respectively (the names referring to the most C‐terminal amino acids in the cytoplasmic tail). In neuromuscular junctions, transmembrane CXADR is exclusively expressed as the SIV isoform (Shaw et al., 2004). The soluble isoforms all lack exon 6 (encoding the transmembrane domain), but contain part of exon 7 (encoding the cytoplasmic tail in the transmembrane isoforms). The soluble isoforms are referred to as CAR4/7, CAR3/7, and CAR2/7, respectively (the nomenclature referring to the exons between which alternative splicing occurs in the individual isoforms) (Do¨rner et al., 2004).

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4.2.4 Posttranslational Modifications CXADR contains two N‐linked glycosylation sites (Tomko et al., 1997). Furthermore, CXADR is palmitoylated at C259 and C260 (van’t Hof and Crystal, 2002).

4.2.5 Homophilic Interactions CXADR has been demonstrated to induce cellular aggregation as a result of homophilic trans‐interactions (Honda et al., 2000; Cohen et al., 2001) (> Table 2-9). However, the regions of the EC modules responsible for the interaction have not been identified.

. Table 2-9 Binding partners for CXADR CAM Igsf11

CXADR CXADR

Extracellular binding partner Igsf11 (trans) AMICA (trans) CXADR (trans) Adenoviruses 2 and 5 Coxsackie viruses B3 and B4 LNX LNX2 MAGI‐1b MPDZ PICK1 PSD‐95 ZO‐1

Reference Harada et al. (2005) Zen et al. (2005) Honda et al. (2000); Cohen et al. (2001) Bergelson et al. (1997) Bergelson et al. (1997) Sollerbrant et al. (2003) Mirza et al. (2005) Excoffon et al. (2004) Coyne et al. (2004) Excoffon et al. (2004) Excoffon et al. (2004) Cohen et al. (2001)

4.2.6 Heterophilic Binding Partners Extracellular Binding Partners CXADR is a receptor for adenoviruses 2 and 5 (which are responsible for respiratory and gastrointestinal infections) and coxsackie viruses B3 and B4 (which cause meningoencephalitis, implicated in acute pancreatitis, and are triggering agents in childhood‐onset diabetes and infections of the heart) (Bergelson et al., 1997). The crystal structure of the CXADR– adenovirus complex has been published, which reveals the interaction of the so‐called knob trimers of the adenovirus with three CXADR D1‐modules (> Table 2-9) (> Figure 2-4) (Bewley et al., 1999). In addition, CXADR forms trans‐interactions with the related molecule AMICA, an interaction reported to involve the membrane‐distal D1‐module of CXADR interacting with the membrane‐proximal Ig‐module of AMICA (Zen et al., 2005). Intracellular Binding Partners In epithelial tissues CXADR localizes to tight junctions. Another tight junction protein, ZO‐1, can be co‐immunoprecipitated with CXADR, possibly owing to interaction with the PDZ‐interacting motif of CXADR (Cohen et al., 2001). Furthermore, CXADR has been demonstrated—through its PDZ‐interacting motif—to bind proteins MAGI‐1b (an epithelial protein, which also interacts with b‐catenin), PICK1 (an epithelial adaptor protein linking transmembrane receptors with PKC), MPDZ (involved in NMDA receptor signaling), PSD‐95 (a palmitoylated neuronal‐ specific protein important for the formation of postsynaptic multiprotein complexes), LNX (an E3 ubiquitin ligase known to mediate ubiquitination and subsequent proteasomal degradation of the


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. Figure 2-4 CXADR. (a): Ribbon diagram of the NMR structure of the first Ig‐module (termed D1) of CXADR. The yellow line indicates the disulfide bridge. (b): Models of the D1 module with colored regions depicting homophilic dimerization (cyan, left), adenovirus binding (Ad) (red, middle), and coxsackie virus binding (CVB) (yellow, right). (c): A schematic representation of the interactions of CXADR in relation to cell adhesion and virus binding (modified from Jiang et al., 2004)

protein NUMB), and the related protein LNX2 (Sollerbrant et al., 2003; Coyne et al., 2004; Excoffon et al., 2004) (> Table 2-9).

4.2.7 Function CXADR acts both as a viral receptor and as an adhesion molecule. Since the receptor predominantly is expressed in the immature neuroepithelium—including various progenitor cells—its main function as an adhesion molecule may be to regulate migration and fasciculation during a restricted period (Hotta et al., 2003). Furthermore, the expression of CXADR on endothelial cells seems to facilitate the transendothelial migration of AMICA‐expressing neutrophils (Zen et al., 2005), whereas it inhibits the migration of cancer cells (Bru¨ning and Runnebaum, 2004). Finally, CXADR is expressed in neuromuscular junctions. This is a feature shared with the adhesion molecules NCAM and N‐cadherin, and like NCAM, CXADR is uniformly expressed on immature muscles, but exclusively restricted to the neuromuscular junctions postnatally (Shaw et al., 2004). This indicates an involvement of CXADR in the organization, development, or maintenance of neuromuscular junctions.

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Studies have shown that disruption of CXADR‐mediated cell–cell interactions increases adenoviral gene transfer, indicating that the localization of CXADR at cell–cell interaction sites and/or homophilic CXADR trans‐interactions restrict/counteract adenoviral binding (Bru¨ning and Runnebaum, 2003). Neither the cytoplasmic tail nor the palmitoylation is important for viral infection (Walters et al., 2001; van’t Hof and Crystal, 2002; Excoffon et al., 2004). However, the cytoplasmic tail is important for the adhesive properties of the molecule. Thus, transepithelial resistance is reduced in cells expressing CXADR receptors lacking the PDZ‐interacting motif. Furthermore, CXADR expression has been shown to reduce the growth of epithelial cells and fibroblasts and tumour cells from human bladder and prostate, and this effect is dependent on both palmitoylation of the cytoplasmic tail and the presence of the PDZ‐binding motif (van’t Hof and Crystal, 2002; Excoffon et al., 2004).

4.3 IGSF11 4.3.1 Gene Locus information for IGSF11 is given in > Table 2-8. IGSF11 was cloned from humans and mice in 2002 by Suzu and coworkers. The gene consists of 10 exons and gives rise to at least two isoforms because of alternative splicing (Katoh and Katoh, 2003).

4.3.2 Expression IGSF11 is predominantly expressed in the brain and testis. In the brain, IGSF11 is preferentially expressed in pyramidal cell layers in the hippocampus and in commissural fibers of the corpus callosum (Suzu et al., 2002). In vitro, the protein is expressed by both glial cells and neurons (Suzu et al., 2002). The expression of the protein is isotype specific. Thus, one isoform in the brain is specifically expressed in the hippocampus (Katoh and Katoh, 2003). Furthermore, IGSF11 is frequently upregulated in different types of cancers, and this expression is also isotype specific (Katoh and Katoh, 2003; Watanebe et al., 2005).

4.3.3 Isoforms and Protein Structure IGSF11 is up to 431‐ and 428 amino acid long in humans and mice, respectively. The transmembrane segment stretches from around amino acid 241–261, followed by a 167–170‐amino acid‐long cytoplasmic tail. The C‐terminal part of the protein contains a sequence (GSLV), which may interact with cytoplasmic scaffolding proteins containing PDZ modules. The human and murine proteins are 88% identical. Furthermore, IGSF11 shows homology to other members of the CTX family (29–30% identity among murine IGFSF11, CXADR, and ESAM) (Suzu et al., 2002; Katoh and Katoh, 2003).

4.3.4 Posttranslational Modifications IGSF11 contains three potential N‐linked glycosylation sites (Suzu et al., 2002).

4.3.5 Homophilic Interactions IGSF11 has been demonstrated to induce cellular aggregation as a result of homophilic trans‐interactions (Harada et al., 2005) (> Table 2-9). However, the regions of the EC modules responsible for the interaction have not been identified.


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4.3.6 Heterophilic Binding Partners The potential heterophilic interactions of IGSF11 with extracellular and intracellular binding partners have to our knowledge not been investigated.

4.3.7 Function IGSF11 is often upregulated in gastric cancers, colon cancers, and hepatocellular carcinomas (Katoh and Katoh, 2003; Watanebe et al., 2005). Transfection of NIH3T3 fibroblasts (which normally do not express this protein) with IGSF11 has been reported to increase cell growth, whereas inhibition of IGSF11 expression in gastric cancer cells by siRNA leads to a reduction in cell growth (Watanebe et al., 2005). However, in a study in which a myeloid leukemia cell line was transfected with IGSF11, the protein had no effect on cell growth (Harada et al., 2005).

5

Neuroplastin

5.1 Introduction Neuroplastin was originally identified as two synaptic glycoproteins with molecular weights of 55 and 65 kDa (Np55 and Np65), respectively (Hill et al., 1988) (> Table 2-10). However, Np55 and Np65 were later shown to be different isoforms of the same protein. The mouse gene was cloned in 1996 (Shirozu et al., 1996), and the human gene was cloned the following year (Langnaese et al., 1997). . Table 2-10 Nomenclature for neuroplastin CAM name Neuroplastin

Protein and gene synonyms Glycoprotein 55, Glycoprotein 65, Np55, Np65, gp55, gp65, stromal cell derived factor receptor 1, SDFRI

5.2 Gene The two isoforms of neuroplastin are the result of alternative splicing of the transcript from a single gene, and in mice and humans, mRNAs of around 2.1 and 2.6 kb have been identified (Shirozu et al., 1996; Langnaese et al., 1997) (> Table 2-11). . Table 2-11 Gene information for neuroplastin Species Man Mouse

Approved gene symbol NPTN Nptn

Approved gene name Neuroplastin

Locus 15q22 9 syntenic

Database reference HGNC ID: 17867 MGI ID: 108077

5.3 Expression Np55 is expressed in a variety of tissues, including lung, liver, and all brain regions. Np65 is brain specific with particularly high expression in forebrain postsynaptic densities where Np55 expression is absent.

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Np65 is also expressed in the midbrain, but not in the brainstem (Langnaese et al., 1997, 1998; Smalla et al., 2000). Np55 is expressed in the embryonic brain and reaches a maximum at around P(10). Np65 is exclusively expressed postnatally, and the expression pattern is proportional to the formation of synapses (Marzban et al., 2003).

5.4 Isoforms and Primary Protein Structure Neuroplastin is a transmembrane protein composed of an extracellular part, followed by a transmembrane region, and a cytoplasmic tail. The two isoforms differ in their extracellular part: Np65 has three Ig‐modules, Np55 only two Ig‐modules (Marzban et al., 2003).

5.5 Posttranslational Modifications Np55 and Np65 are glycoproteins, and both isoforms contain six potential sites for N‐linked glycosylation. Deglycosylation experiments have revealed that both isoforms contain high mannose levels and complex oligosaccharides. The glycosylation patterns have been shown to vary with the differential expression of the proteins in different cell types (Langnaese et al., 1998). Np65 has also been shown to contain fucose, which may play an important role in long‐term potentiation (LTP) (Smalla et al., 1998, 2000).

5.6 Homophilic Interactions Only the Np65 isoform of neuroplastin is engaged in homophilic binding, which suggests that the Np65‐ specific Ig‐module plays an important role in this interaction. In concordance with this, it has been demonstrated that constructs containing Ig‐module 1–3, but not constructs containing only Ig‐module 2–3, mediate trans‐homophilic binding (Smalla et al., 2000) (> Table 2-12).

. Table 2-12 Binding partners for neuroplastin CAM Np65

Extracellular binding partner Np65 (trans)

Reference Smalla et al. (2000)

5.7 Function The most prominent function of neuroplastin recognized so far is cell adhesion. It is believed that Np65 binds homophilically in trans at the synaptic cleft, and thus contributes to synaptogenesis and synapse stabilization. Neuroplastins are also involved in hippocampal LTP. Thus, anti-neuroplastin antibodies block LTP in hippocampal slices. Likewise, Np65 levels in the postsynaptic density are upregulated after LTP induction (Smalla et al., 2000).

6

Signal‐Regulatory Proteins

6.1 Introduction The family of ‘‘signal regulatory proteins’’ (SIRPs) has been reported to include at least 15 members (Kharitonenkov et al., 1997). However, so far only a few genes have been cloned. The orthologous genes for


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PTPNS1 (protein tyrosine phosphatase, non‐receptor type substrate 1) in humans, rats, and mice were cloned in 1996 and have since been cloned multiple times, and consequently, the protein has been given many names including SHPS‐1 (SHP substrate 1) (Fujioka et al., 1996; Yamao et al., 1997), SIRPa1 (Kharitonenkov et al., 1997), BIT (brain immunoglobulin‐like molecule with tyrosine‐based activation motifs) (Ohnishi et al., 1996, 1997; Sano et al., 1997), and MFR (macrophage fusion receptor) (Saginario et al., 1998). Before the cloning of PTPNS1, the protein was also referred to as gp93 (glycoprotein 93) (Li et al., 1992; Quiroga and Pfenninger, 1994; Wang et al., 2003) and p84 (Comu et al., 1997). In the following sections the protein is referred to as PTPNS1 (> Table 2-13). . Table 2-13 Nomenclature for SIRP family CAM name PTPNS1 PTPNS1L PTPNS1L2 PTPNS1L3 SIRPB1 SIRPB2

Protein and gene synonyms CD172a, SHPS1, SIRP, gp93, P84, MYD‐1, BIT, SHPS‐1, SIRPa, SIRPa2, MFR, SIRPa‐1 dJ576H24.4 dJ776F14, PTPN1L SIRP‐BETA‐1, 9930027N05Rik bA77C3.1, SIRPg

Besides PTPNS1, the SIRP family also includes SIRP‐B1/SIRPb and SIRP‐B2/SIRPg (Kharitonenkov et al., 1997; Adams et al., 1998; Ichigotani et al., 2000; Brooke et al., 2004). These proteins are predominantly found in monocytes, dendritic cells, T cells, and B cells, and therefore not described in detail here. Finally, three PTPNS1‐like proteins have been cloned from humans (PTPNS1L, PTPNS1L2, PTPNS1L3). However, these proteins have not been investigated further, and therefore not described here (> Table 2-13 and > 2-14).

. Table 2-14 Gene information for SIRP family members

Species Man

Approved gene symbol PTPNS1 PTPNS1L PTPNS1L2 PTPNS1L3

Mouse

SIRPB1 SIRPB2 Ptpns1 Sirpb

Approved gene name Protein tyrosine phosphatase, nonreceptor type substrate 1 Protein tyrosine phosphatase, nonreceptor type substrate 1‐like Protein tyrosine phosphatase, nonreceptor type substrate 1‐like 2 Protein tyrosine phosphatase, nonreceptor type substrate 1‐like 3 Signal‐regulatory protein b1 Signal‐regulatory protein b2 Protein tyrosine phosphatase, nonreceptor type substrate 1 Signal‐regulatory protein b (orthologue to human SIRPB1)

Locus 20p13


22q12.2

HGNC ID: 9663

20

HGNC ID: 16248

20p12.2–13

HGNC ID: 16247

20 20p13 2 73.1 F3

HGNC ID: 15928 HGNC ID: 15757 MGI ID: 108563

3 syntenic

MGI ID: 2444824

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6.2 Genes Human PTPNS1 mRNA exists in at least two isoforms, indicating that the gene product is the subject of alternative splicing (Yamao et al., 1997). In mouse, the Ptpns1 consists of eight exons. Exon 1 encodes a signal peptide. Exons 2, 3, and 4 each encode an Ig‐module. Exon 5 encodes a transmembrane segment, and the last three exons encode the cytoplasmic tail of the protein (Sano et al., 1999). Like the human protein, murine PTPNS1 exists in several isoforms as a result of alternative splicing (Comu et al., 1997; Veillette et al., 1998; Sano et al., 1999). Thus, exon 7 has two splice donor sites, resulting in the production of mRNAs, which do or do not contain a sequence encoding a VQSL sequence in the cytoplasmic tail of the proteins. In addition, one splice variant does not include exon 3 (the middle Ig‐module), another splice variant does not include any of the exons encoding the extracellular Ig‐modules (Sano et al., 1999), and another that extracellularly only contains the first (membrane‐distal) Ig‐module has also been reported (Comu et al., 1997).

6.3 Expression PTPNS1 is expressed in most tissues. In humans, the protein is predominantly expressed in the brain, but found also in the thymus, intestine, lung, spleen, and leukocytes. In rat, the protein is predominantly expressed in the brain, spleen, and lung (Fujioka et al., 1996; Sano et al., 1997; Yamao et al., 1997). The protein is abundant in macrophages, but—in contrast to SIRP‐B2—not expressed in T cells and B cells (Veillette et al., 1998). In mouse, PTPNS1 is present already at E7 (Yamao et al., 1997), and at E9 it can be detected in the floor plate region of the embryonic spinal cord (Comu et al., 1997). Later, it is expressed in the developing retina, where it is present in both the inner and the outer plexiform layers and in photoreceptor cells (Mi et al., 2000). Postnatally, the protein is more widely expressed in the central nervous system (CNS), including cerebellum, cerebral cortex, hippocampus, olfactory bulb, and spinal cord (Comu et al., 1997). In rats, where the distribution of PTPNS1 is determined at E15, E18, and P3, the protein is in general most abundant in the early embryonic stages. In the nervous system the protein is expressed in the neuroepithelium, cerebral cortex, septo‐hippocampal pathway, brainstem, and midbrain (Li et al., 1992; Henry et al., 1999), but it is also found in macrophages, monocytes, granulocytes, and dendritic cells (Adams et al., 1998).

6.4 Isoforms and Protein Structure Human, rat, and mouse PTPNS1 proteins are up to 503‐, 509‐, and 509 amino acid long, respectively, and the sequences of the human, mouse, and rat proteins are 65–81% identical and >91% similar (Yamao et al., 1997). The protein contains a 28‐amino‐acid signal peptide, three Ig‐modules (the membrane‐distal module is of the V‐type; the other two modules are of the C1‐type), a transmembrane segment (amino acid 344–368 of mouse PTPNS1), and a cytoplasmic tail. The cytoplasmic tail contains four tyrosine residues (Y408, Y432, Y449, and Y473 of mouse PTPNS1). These residues are positioned in a consensus motif (YXX (L/V/I)) that represents potential tyrosine phosphorylation sites, referred to as tyrosine‐based activation motifs (TAMs). The four TAMs (YADL, YASI, YADL, and YASV) can, if phosphorylated, bind Src homology 2 (SH2)‐modules and are completely conserved between humans, mice, and rats (Comu et al., 1997; Yamao et al., 1997). In addition to the various isoforms derived from the alternative splicing mentioned above, PTPNS1 also exists in a soluble, shed form derived from the cleavage of the molecule between S359 and M360, close to the transmembrane segment. The cleavage is believed to be mediated by metalloproteinases (probably MMP‐1 and ‐9) (Ohnishi et al., 2004).


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6.5 Posttranslational Modifications In addition to the phosphorylation of the tyrosine residues in the above‐mentioned TAMs, PTPNS1 also contains several potential serine/threonine‐phosphorylation sites (Fujioka et al., 1996), but the importance of these for the function of SIRPa is not known. Extracellularly, the protein contains several potential N‐glycosylation sites (17 in mouse, 15 in rat, and 4 in human) (Fujioka et al., 1996; Comu et al., 1997; Yamao et al., 1997). The protein is usually highly glycosylated, and binds to concanavalin A (ConA), wheat germ agglutinin (WGA), and Galanthus nivalis agglutinin (GNA). The main type of glycosylation is Mana1‐6(Mana1‐3)Mana1‐6(Mana1‐3)‐Manb1‐ 4GlcNAcb1‐4GlcNAc, whereas the protein apparently does not contain any HNK‐1 epitopes (an epitope expressed on a number of CAMs, including NCAM, L1, TAG‐1, and P0) (Li et al., 1992; Quiroga and Pfenninger, 1994; Fujioka et al., 1996; Bartoszewicz et al., 1999).

6.6 Homophilic Interactions Members of the SIRP family have not been reported to form homophilic interactions.

6.7 Heterophilic Binding Partners 6.7.1 Extracellular Binding Partners PTPNS1 has in several studies been shown to interact in trans with CD47 (described in Sect. 5) (> Table 2-6) (Jiang et al., 1999; Seiffert et al., 1999; Vernon‐Wilson et al., 2000). PTPNS1 binds to CD47 with a Kd of 2–8 mM, and the interactions take place between the membrane‐distal V‐type Ig‐module of PTPNS1 and the single V‐type Ig‐module of CD47 (Han et al., 2000; Vernon‐Wilson et al., 2000; Liu et al., 2002; Brooke et al., 2004). A peptide (CERVIGTGWVRC), which mimics an epitope on CD47, binds in a dose‐dependent manner to the V‐type Ig‐module of PTPNS1. This peptide specifically blocks CD47‐PTPNS1 interactions, and inhibits transmigration of neutrophils across intestinal epithelium in a dose‐dependent manner (Liu et al., 2004a). PTPNS1 has also been reported to interact directly with fibronectin (Oshima et al., 2002a), and the plating of cells on laminin or fibronectin leads to tyrosine phosphorylation of PTPNS1 and subsequent binding of SHP‐2 (see below) (Tsuda et al., 1998).

6.7.2 Intracellular Binding Partners The cytoplasmic tail of PTPNS1 binds to two proteins termed SH2‐containing tyrosine phosphatase‐1 and ‐2, SHP‐1 and ‐2 (also known as HCP, HCPH, PTP1C and SH‐PTP2, SH‐PTP3, PTP1D, PTP2C, SAP‐2, and Syp, respectively) (> Table 2-15) (Fujioka et al., 1996; Ohnishi et al., 1996; Kharitonenkov et al., 1997). SHP‐1 and ‐2 are nonreceptor protein tyrosine phosphatases located in the cytosol. They are composed of two SH2 modules followed by a classical protein tyrosine phosphatase module and a C‐terminal tail (reviewed by Neel et al., 2003). The interaction between PTPNS1 and SHP‐1 and ‐2 takes place between the phosphorylated TAMs of PTPNS1 and the SH2 modules of SHP‐1 and ‐2. Thus, SHP‐1 and‐2 proteins lacking their SH2 modules do not bind to PTPNS1, and PTPNS1, in which the tyrosine residues of the TAMs have been replaced with phenylalanine, does not bind SHP‐1 or ‐2 (Fujioka et al., 1996; Araki et al., 2000a). Studies have demonstrated that the tandem SH2 modules bind distinct TAMs with a Kd around 0.5–3.0 nM, whereas the affinities between tandem SH2 modules and TAMs that are not natural biological partners are 100‐ to 10,000‐fold lower. The Kd between PTPNS1 and SHP‐2 is 1.3 nM (Ottinger et al., 1998). The tyrosine residues on PTPNS1 responsible for binding to SHP‐2 have been reported to be Y449 and Y473 (Takada et al., 1998).

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. Table 2-15 Binding partners for PTPNS1 Extracellular binding partners CD47 (trans) Fibronectin SHP‐1, SHP‐2 Grb2 JAK2 PYK2 SKAP55hom/R CSF‐1R

Reference Jiang et al. (1999); Seiffert et al. (1999); Vernon‐Wilson et al. (2000); Han et al. (2000) Oshima et al. (2002) Fujioka et al. (1996); Ohnishi et al. (1996); Kharitonenkov et al. (1997); Takada et al. (1998) Kharitonenkov et al. (1997) Stofega et al. (1998) Timms et al. (1999) Timms et al. (1999) Timms et al. (1998)

PTPNS1 is also reported to bind to the adaptor protein Grb2 and the kinase JAK2 (Kharitonenkov et al., 1997; Stofega et al., 1998). Furthermore, the protein has in macrophages been reported to bind to the tyrosine kinase PYK2, the adaptor protein SKAP55hom/R, and the tyrosine kinase receptor CSF‐1R. However, the interaction sites have not been identified (> Table 2-15) (Timms et al., 1998, 1999).

6.8 Signaling PTPNS1 is a substrate for several enzymes including the growth factor receptors PDGFR, EGFR, and IR, and the cytoplasmic tyrosine kinase Src and SHP‐2 (Fujioka et al., 1996; Kharitonenkov et al., 1997; Tsuda et al., 1998). The TAM‐tyrosine residues of PTPNS1 are rapidly phosphorylated in response to treatment of cells with the growth factors EGF, GH, HGF/SF, NGF, BDNF, neurotrophic factor NT‐3, and insulin, the lipid lysophosphatidic acid (LPA), which activates various G‐protein‐coupled receptors, and serum (Fujioka et al., 1996; Ochi et al., 1997; Ohnishi et al., 1999; Yan et al., 2004). In addition, adhesion of cells to fibronectin also induces a marked increase in tyrosine phosphorylation of PTPNS1 (Fujioka et al., 1996). Subsequent to phosphorylation of the TAM tyrosine residues, PTPNS1 binds to SHP‐1 and/or SHP‐2 (Fujioka et al., 1996; Yan et al., 2004). This interaction can stimulate the protein phosphatase activity of SHP. Thus, a peptide corresponding to the TAM‐region of PTPNS1 induces the enzyme activity of SHP‐2 in vitro (Ohnishi et al., 1996). The response of PTPNS1 to growth factor stimulation is cell dependent. Thus, NGF stimulates PTPNS1‐phosphorylation and SHP‐2 binding in PC12‐cells, but not in primary cultures of cerebral cortical neurons (Ohnishi et al., 1999). Tyrosine phosphorylation and overexpression of PTPNS1 has been reported to affect DNA synthesis and the activity of MAPKs. However, the reported observations are not consistent. In some studies MAPK activity was inhibited, whereas other studies report activation (Kharitonenkov et al., 1997; Takada et al., 1998; Tsuda et al., 1998; Yan et al., 2004), and in some cases, overexpression of PTPNS1 had no effect on MAPK activity (Ochi et al., 1997; Araki et al., 2000b; Wu et al., 2000). Induction of tyrosine phosphorylation of PTPNS1 and the subsequent association with SHP‐2 and MAPK activation can be inhibited by the Clostridium botulinum exoenzyme C3, an inhibitor of the GTPase Rho, and it is also inhibited in FAK‐deficient cells, suggesting that the process is mediated by a Rho‐ dependent pathway that includes FAK and a Src family kinase (Takeda et al., 1998). Interestingly, treatment of cells with antibodies against PTPNS1 can enhance the activity of Rho and the subsequent formation of


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actin stress fibers (Motegi et al., 2003), indicating that Rho may be located both upstream and downstream of PTPNS1‐mediated signaling. In neuroblastoma cells, the extracellular part of PTPNS1 can induce CD47‐ mediated trans‐activation of two other small GTPases, Rac and Cdc42, subsequently leading to the formation of neurites and filopodia, respectively (Miyashita et al., 2004). Overexpression of PTPNS1 has also been reported to reduce EGF‐induced PI3K activity, possibly by reducing the interaction between SHP‐2 and the regulatory p85 subunit of PI3K (Wu et al., 2000), whereas overexpression in neurons increases BDNF‐induced activation of the protein kinase PKB in a manner that appears to be independent of tyrosine phosphorylation of PTPNS1, but dependent on PI3K activity (Araki et al., 2000a). Finally, overexpression of PTPNS1 is also reported to reduce activation of NF‐kB (Yan et al., 2004), a transcription factor involved in the regulation of immune responses, cell growth, differentiation, and proliferation.

6.9 Function Although PTPNS1 is not involved in homophilic trans‐interactions, one of its functions clearly is to mediate intercellular adhesion. Thus, cells can adhere to a substrate of immobilized PTPNS1 within 10 min of plating (Chuang and Lagenaur, 1990). The main trans‐interacting partner of PTPNS1 is most likely CD47, and indeed the distribution of the two proteins is almost identical in the adult brain (Jiang et al., 1999). Neurotrophins such as NGF and BDNF, which can induce tyrosine phosphorylation of PTPNS1, play a role in cellular survival, differentiation, and synaptic plasticity in the nervous system (Ohnishi et al., 1999). Consistently, BDNF‐promoted survival of cultured cerebral cortical neurons can be improved by overexpression of PTPNS1, and as mentioned above, this effect is not dependent on the tyrosine phosphorylation of PTPNS1 (Araki et al., 2000a). PTPNS1‐coated substrates have been demonstrated to promote neurite outgrowth (Sano et al., 1997; Miyashita et al., 2004). Neurites and growth cones contain PTPNS1 (Quiroga and Pfenninger, 1994), and axons of brain tissue transplants express large amounts of PTPNS1, indicating that the molecule is important for neural development and regeneration (Henry et al., 1999). Furthermore, the protein is upregulated in denervated muscles, indicating that it may be involved in interactions between neurons and muscle cells (Mitsuhashi et al., 2005). The morphology of neurites for cells grown on immobilized PTPNS1, NCAM, or L1 differs, indicating that these molecules promote neurite outgrowth by distinct mechanisms (Abosch and Lagenaur, 1993). Recent studies indicate that PTPNS1 may be involved in the maintenance of homeostasis of various physiological conditions. Thus, tyrosine phosphorylation of PTPNS1 is enhanced in selected regions of the hypothalamus in response to exposure of rats to hypothermia (Taniguchi et al., 2004). Furthermore, several studies suggest a role of PTPNS1 in relation to regulation of the circadian clock. Thus, tyrosine phosphorylation of PTPNS1 in the hypothalamic suprachiasmatic nucleus and the retina is higher in light periods than in dark periods, and it is increased after light exposure in dark periods. The phosphorylation leads to recruitment of SHP‐2 to PTPNS1 and is in part dependent on the function of NMDA receptors (Nakahata et al., 2000, 2003; Hamada et al., 2004). Expression of PTPNS1 is affected in certain types of cancer. Thus, the expression is found to be lower in certain hepatocellular carcinoma and breast cancer cells than in corresponding normal tissues. Furthermore, PTPNS1 is often downregulated in cells transformed by v‐Src, k‐Ras, v‐Fps, and papillomavirus large T (Oshima et al., 2002), whereas overexpression of PTPNS1 in carcinoma and glioblastoma cells can suppress tumor growth in vitro and in vivo and reduce cellular migration and cell spreading (Wu et al., 2000; Yan et al., 2004). These effects seem to be dependent on the tyrosine phosphorylation of PTPNS1. Expression of mutant versions of PTPNS1, which either cannot be shed (because of mutations in the membrane‐proximal extracellular part of the protein) or cannot be tyrosine phosphorylated (because of deletion of the cytoplasmic tail), leads to a more rapid adhesion of the cells to a fibronectin substrate, followed by increased formation of actin stress fibers and focal adhesions, increased phosphorylation of FAK and paxillin, and impairment of cell migration, indicating that the protein regulates the dynamics of the actin cytoskeleton (Inagaki et al., 2000; Ohnishi et al., 2004).

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Macrophages express CD47 as well as PTPNS1, and abrogation of CD47–PTPNS1 interactions prevents macrophage fusion (Saginario et al., 1998; Han et al., 2000). In these cells, PTPNS1 predominantly binds to SHP‐1 rather than SHP‐2 (Veillette et al., 1998). PTPNS1–CD47 interactions inhibit the spreading and aggregation of platelets, an effect which in part involves the inhibition of integrin aIIbb3‐mediated outside‐in signaling (Kato et al., 2005). The PTPNS1–CD47 interaction is also important for chemotaxis and transmigration of neutrophils (Liu et al., 2002), and for interactions between T cells and myelomonocytes (Latour et al., 2001). Interaction between PTPNS1 and CD47 is also important for the transmigration of monocytes across the cerebral endothelium, a process of central importance for the development of neuroinflammatory diseases (de Vries et al., 2002). In contrast, attachment of monocytes to the elastic laminae of arteries seems to be inhibited by PTPNS1 (Liu et al., 2005). For reviews of PTPNS1 see Cant and Ullrich (2001) and Oshima et al. (2002b).

7

Nectins (Poliovirus Receptor‐Related Proteins)

7.1 Introduction Nectins constitute a family of four proteins (nectin‐1 to ‐4); (> Table 2-16). They share homology to the poliovirus receptor (necl‐5) and are therefore also called poliovirus receptor‐related 1–4, PRR1–4. Like the . Table 2-16 Nomenclature for nectin family CAM name Nectin‐1 Nectin‐2 Nectin‐3 Nectin‐4

Protein and gene synonyms CD111, CLPED1, ED4, HGNC:3160, HIgR, HVEC, poliovirus receptor‐related 1, PRR, PRR1, PVRR1, SK‐12 CD112, HVEB, MPH, poliovirus receptor‐related 2, PRR2, Pvr, PVRR2, Pvs 2610301B19Rik, 3000002N23Rik, 4921513D19Rik, DKFZP566B0846, poliovirus receptor‐related 3, PPR3, PVRR3 1200017F15Rik, LNIR, poliovirus receptor‐related 4, PRR4

poliovirus receptor these proteins serve as receptors for various viruses. Thus, nectin‐1 mediates entry of herpes simplex viruses HSV‐1 and HSV‐2, PRV (porcine pseudorabies virus), and BHV‐2 (bovine herpesvirus) (Geraghty et al., 1998), whereas nectin‐2 mediates entry of HSV‐2, various mutant strains of HSV‐1, and pseudorabies virus (Warner et al., 1998). Consequently, nectin‐1 and ‐2 are also known as herpes virus entry mediator C and B, respectively. In addition to their virus‐binding capabilities, nectins are CAMs (hence the name nectin; from Latin ‘‘necto’’ meaning ‘‘to connect’’) (Takahashi et al., 1999), and the following sections will focus on nectins as CAMs. For a review of the virus‐related properties of the nectins, see Campadelli‐Fiume et al. (2000). The poliovirus receptor and four related proteins constitute a separate family, the nectin‐like proteins, necls. The general structure of nectins and necls is the same, and the distinction between the two families is based solely on the ability of the proteins to bind to the cytosolic protein afadin. Thus, nectins bind to afadin, whereas necls supposedly do not (> Figure 2-5) (see also > Sect. 10) (Takai et al., 2003b).

7.2 Genes The gene encoding murine nectin‐2 was cloned in 1992 (Morrison and Racaniello, 1992; Aoki et al., 1994, 1997) followed by the genes encoding human nectins‐1 and ‐2 (Eberle´ et al., 1995; Lopez et al., 1995; Cocchi et al., 1998), human nectins‐3 and ‐4, and mouse nectins‐1, ‐3, and ‐4 (Menotti et al., 2000; Reymond et al., 2000, 2001; Satoh‐Horikawa et al., 2000; Lopez et al., 2001) (> Table 2-17).


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. Figure 2-5 Nectins and Necls. Organization of (a) nectins, (b) afadin, (c) afadin‐binding proteins, and (d) Necls. Nectins bind afadin via the PDZ‐domain binding sequence E/AxYV (red in (a)). This sequence is absent in Necls (d). (c) Afadin can via different domains interact with several other proteins. CC, coiled‐coil domain; CH calponin‐homology domain; DIL, dilute domain; FHA, forkhead‐associated domain; LIM, LIM domain; PDZ, PDZ domain; PR, proline‐ rich domain; RA, Ras‐associated domain; SH3, Src‐homology 3 domain; TM, transmembrane region (modified from Irie et al., 2004)

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. Table 2-17 Gene information for nectin family members

Species Man

Approved gene symbol PVRL1 PVRL2

Mouse

PVRL3 PVRL4 Pvrl1 Pvrl2 Pvrl3 Pvrl4

Approved gene name Poliovirus receptor‐related 1 (herpesvirus entry mediator C; nectin) Poliovirus receptor‐related 2 (herpesvirus entry mediator B) Poliovirus receptor‐related 3 Poliovirus receptor‐related 4 Poliovirus receptor‐related 1 Poliovirus receptor‐related 2 Poliovirus receptor‐related 3 Poliovirus receptor‐related 4

Locus 11q23–q24


19q13.2–q13.4

HGNC ID: 9707

3q13 1q22–q23.2 9B 7 9.0 16 syntenic 1 H2

HGNC ID: 17664 HGNC ID: 19688 MGI ID: 1926483 MGI ID: 97822 MGI ID: 1930171 MGI ID: 1918990

7.3 Expression Nectin‐1 is widely expressed in the nervous system of both humans and mice (Cocchi et al., 1998; Haarr et al., 2001) including the CA3 area of the hippocampus, where it colocalizes with the cytosolic proteins afadin and ZO‐1 (Nishioka et al., 2000; Inagaki et al., 2003). In humans nectin‐3 is mainly expressed in testis and placental tissues (Reymond et al., 2000), whereas in mice it is also expressed in the neuroepithelium (Okabe et al., 2004). In the adult mouse hippocampus, nectin‐1 and ‐3 are both present at puncta adherentia and at synaptic junctions between mossy fibre terminals and dendrites of pyramidal cells where they are localized on pre‐ and postsynaptic membranes, respectively (Mizoguchi et al., 2002; Yamada et al., 2003). Nectin‐2 is ubiquitously expressed, mainly in general columnar epithelia, but is also expressed in ovarian granulosa cells during follicular development (Okabe et al., 2004; Kawagishi et al., 2005). Nectin‐4 is in human tissues mainly restricted to the placenta, whereas the expression in mouse is broader, including brain, lung, and testis (Reymond et al., 2001).

7.4 Isoforms and Protein Structure Nectins are composed of a 324‐amino acid‐long extracellular part followed by a single transmembrane segment and a 140‐amino acid‐long cytoplasmic tail. The extracellular parts of the molecules consist of three Ig‐modules. The N‐terminal module is of the V‐type, the second of the C1‐type, and the third module of the C2‐type (Du Pasquier et al., 2004). Each of the four nectins exists in up to three isoforms, a, b, and g, because of alternative splicing of the transcripts. The three isoforms of nectin‐3 are transmembrane proteins, which mainly differ in the length of the cytoplasmic tail (nectin‐3a being the longest, and ‐3g the shortest isoform) (Satoh‐Horikawa et al., 2000). However, at least one nectin, nectin‐1g, is a secreted isoform, which does not contain the transmembrane and cytoplasmic regions. Moreover, of the nectin isoforms identified so far, nectins‐1a, ‐2a, ‐2d, ‐3a, and 3b contain the C‐terminal sequence (A/E)‐x‐Y‐V, which is a consensus sequence mediating interaction with the PDZ domain of afadin, whereas nectins‐1b, ‐3g and ‐4 do not contain this sequence (Reymond et al., 2001).

7.5 Posttranslational Modifications Nectins contain up to eight potential N‐glycosylation sites. Studies indicate that most or all of these glycosylation sites can be glycosylated (Krummenacher et al., 1998).


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Nectin‐2d becomes phosphorylated on tyrosine residues in response to trans‐homophilic dimerization (Lopez et al., 1998). The mouse isoform of nectin‐2d contains four potential tyrosine phosphorylation sites (Y505, Y510, Y517, and Y529) of which Y505 appears to be the predominant site of phosphorylation (Kikyo et al., 2000).

7.6 Nectin–Nectin Interactions All nectins form Ca2þ/Mg2þ‐independent dimers by homophilic cis‐interactions (Mizoguchi et al., 2002). However, with the exception of nectin‐2a, which can dimerize with nectin‐2d (Lopez et al., 1998), they appear not to form heterophilic cis‐dimers. Furthermore, all nectins are involved in homophilic and/or heterophilic trans‐interactions. Thus, nectin‐1 interacts in trans with nectins‐3 and ‐4, and nectin‐3 interacts in trans with nectins‐2 and ‐3 (Satoh‐Horikawa et al., 2000; Reymond et al., 2001; Fabre et al., 2002) (> Table 2-18). The heterophilic trans‐interactions are reported to have lower Kd‐values than that of the homophilic interactions (Satoh‐Horikawa et al., 2000; Fabre et al., 2002; Yasumi et al., 2003).

. Table 2-18 Binding partners for nectin family members CAM Nectin‐1

Extracellular binding partner Viruses (HSV‐1, HSV‐2, PVR, BHV‐2) Nectin‐1 (cis) Nectin‐3, ‐4 (trans)

Nectin‐2

Viruses (HSV‐1, HSV‐2, pseudorabies virus) Nectin‐2 (cis)

Nectin‐1 (trans) Afadin PICK‐1

Reymond et al. (2005)

PAR‐3 Presenilin‐1

Takekuni et al. (2003) Kim et al. (2002)

CD44 Nectin‐3 (cis) Nectin‐1, ‐2 (trans) Necl‐5, SynCAM

Nectin‐4

Nectin‐1a, ‐2a, ‐2d, 3a, ‐3b, and ‐4 Nectin‐1a, ‐2a, ‐2d, 3a, ‐3b, and ‐4 Nectin‐1, ‐3 Nectin‐1

Reymond et al. (2001); Mizoguchi et al. (2002); Yasumi et al. (2003) Fabre et al. (2002); Reymond et al. (2001); Satoh‐Horikawa et al. (2000) Warner et al. (1998) Lopez et al. (1998); Reymond et al. (2001); Mizoguchi et al. (2002); Yasumi et al. (2003) Satoh‐Horikawa et al. (2000); Momose et al. (2002) Bottino et al. (2003); Tahara‐Hanaoka et al. (2004); Pende et al. (2005) Ref203Freistadt and Eberle, (1997) Reymond et al. (2001); Mizoguchi et al. (2002); Yasumi et al. (2003) Fabre et al. (2002); Satoh‐Horikawa et al. (2000); Momose et al. (2002) Fabre et al. (2002); Ikeda et al. (2003); Mueller and Wimmer (2003); Shingai et al. (2003) Reymond et al. (2001); Mizoguchi et al. (2002); Yasumi et al. (2003) Fabre et al. (2002); Reymond et al. (2001) Mandai et al. (1997)

Nectin‐3 (trans) DNAM‐1

Nectin‐3

Reference Geraghty et al. (1998)

Nectin‐4 (cis)

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The first Ig‐module of nectin‐1 is reported to be involved in the trans‐dimerization with nectins‐3 and ‐4 (Fabre et al., 2002; Momose et al., 2002), whereas the second Ig‐module of the nectins is believed to be important for cis‐dimerization (Reymond et al., 2001; Yasumi et al., 2003). Furthermore, a recombinant construct consisting of all three Ig‐modules of nectin‐1 has been reported to form tetramers, whereas constructs lacking the third Ig‐module only form dimers, indicating that the third Ig‐module also is involved in dimerizations (Krummenacher et al., 1999). Mutation of F136 in the first Ig‐module of mouse nectin‐2 abrogates trans‐interactions but not cis‐interactions (Miyahara et al., 2000), whereas deletion of the Ig2‐module from nectins abrogates not only cis‐ but also trans‐interactions (Momose et al., 2002). Thus, cis‐interactions between nectins are believed to be a prerequisite for subsequent trans‐interactions.

7.7 Heterophilic Binding Partners 7.7.1 Extracellular Binding Partners Nectin‐3 can bind to SynCAM and Necl‐5 (see also Sect. 10 and > Table 2-21); (Fabre et al., 2002; Ikeda et al., 2003; Mueller and Wimmer, 2003; Shingai et al., 2003), and nectin‐2 can bind to DNAM‐1, a leucocyte adhesion molecule expressed on NK cells, T cells, and monocytes (Bottino et al., 2003; Tahara‐Hanaoka et al., 2004; Pende et al., 2005). In monocytes, nectin‐2 has also been reported to form a cis‐association with the receptor for hyaluronic acid, CD44 (Freistadt and Eberle, 1997) (> Table 2-18).

7.7.2 Intracellular Binding Partners Of the nectin isoforms identified so far, nectins‐1a, ‐2a, ‐2d, 3a, ‐3b, and ‐4, but not nectins‐1b and ‐3g, have been demonstrated to bind directly to the cytosolic protein afadin. Afadin exists in two isoforms, a large form termed l‐afadin, which is ubiquitously expressed, and a small form, s‐afadin, which predominantly is expressed in the brain. L‐afadin, but not s‐afadin, contains an F‐actin‐binding domain, and both isoforms contain a PDZ domain (Mandai et al., 1997). All afadin‐interacting nectins—with the exception of nectin‐4—bind to the PDZ domain of afadin via the above‐mentioned consensus sequence, (A/E)‐X‐Y‐V, at their C‐terminal (> Figure 2-5); (Takahashi et al., 1999; Reymond et al., 2000, 2001; Satoh‐Horikawa et al., 2000). Nectins are associated with a large number of cytosolic proteins through their association with afadin. For instance, afadin binds (directly or indirectly) to Eph receptor tyrosine kinases (Buchert et al., 1999), the insulin receptor‐interacting protein ponsin (Mandai et al., 1999), the a‐actinin‐binding protein LMO7 (Ooshio et al., 2004), the F‐actin binding proteins IQGAP1 and ZO‐1 (Yokoyama et al., 2001; Katata et al., 2003), the CTX‐family adhesion molecule F11R (Fukuhara et al., 2002b) (see also > Sect. 6), the a‐actinin‐ binding protein ADIP (Asada et al., 2003), the membrane associated scaffold protein mLin‐7 (Yamamoto et al., 2002), the PDZ domain protein S‐SCAM (Yamada et al., 2003), the scaffold protein PICK‐1 (Reymond et al., 2005), and the multitransmembrane protein presenilin‐1, PS‐1 (Kim et al., 2002). Noticeably, l‐afadin binds to a‐catenin (Tachibana et al., 2000; Pokutta et al., 2002). a‐catenin interacts with the cytoplasmic tail of adhesion molecules belonging to the cadherin family, and therefore the afadin– a‐catenin interaction serves to link nectins with cadherins. The C‐terminal amino acids of nectins‐1 and ‐3 have been reported to interact directly with PAR‐3 (an adaptor protein involved in cell polarization processes and formation of tight junctions), which like afadin binds the nectins through a PDZ domain (Takekuni et al., 2003) (> Table 2-18).

7.8 Signaling Binding of afadin to nectin is believed to facilitate nectin cis‐dimerization and the subsequent trans‐ interactions and clustering with cadherins (Takahashi et al., 1999; Miyahara et al. 2000). For nectin‐2d the


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dimerization leads to phosphorylation of tyrosine residue(s) in the cytoplasmic tail of the molecule (see above) (Lopez et al., 1998). Expression of E‐cadherin promotes this phosphorylation, whereas abrogation of extracellular cadherin interactions reduces the phosphorylation (Kikyo et al., 2000), stressing the importance of functional interactions between nectins and the cadherins. Trans‐interactions of nectins induce the formation of filopodia and lamellipodia through the activation of the GTPases Cdc42 and Rac, respectively. The nectin‐induced activation of Cdc42 and Rac is independent of nectin–afadin interactions, but requires the presence of the cytoplasmic tail of the nectins (Kawakatsu et al., 2002; Fukuhara et al., 2003). Trans‐interacting E‐cadherins also activate Rac and in both cases the increased GTPase activity affects the rate of formation of tight junctions and adherence junctions (Hoshino et al., 2004). Activation of the GTPases ultimately leads to the activation of the MAPK, JNK, but not p38 or ERK (Honda et al., 2003b). Addition of the phorbol ester TPA (which induces activation of PKC) to MDCK cells leads to the formation of tight junction‐like structures with accumulation of nectin and afadin but not cadherin (Asakura et al., 1999). However, treatment of various cell types with TPA or HGF/SF also induces an extracellular proteolytic cleavage of nectin‐1a—but not nectins‐2a or ‐2d—by MMPs. The consequence of proteolysis is the shedding of an 80‐kDa extracellular fragment of nectin, leaving a 26–33‐kDa nectin fragment containing the transmembrane and cytoplasmic tail of the protein in the plasma membrane (Kim et al., 2002; Tanaka et al., 2002). Presenilin‐1, PS‐1, is a subunit of the g‐secretase complex and binds to the C‐terminal part of nectin‐1, and the degradation of the membrane‐bound C‐terminal fragment of nectin can be inhibited by inhibitors of g‐secretases and by expression of dominant negative PS‐1 (Kim et al., 2002).

7.9 Functions Because of their ability to interact in cis‐ and trans, nectins promote aggregation of cells (Aoki et al., 1997; Lopez et al., 1998). In epithelial cells, nectins are important for the formation of tight junctions and adherence junctions (Fukuhara et al., 2002a), and overexpression of nectin‐1 increases the rate of formation of these structures (Honda et al., 2003a). In the nervous system, nectins‐1 and ‐3 are important for the formation of synapses (see > Figure 2-6). Thus, in vitro studies with hippocampal neurons have revealed that inhibition of trans‐interactions between nectins‐1 and ‐3 (by addition of the nectin‐1‐binding protein glycoprotein D, derived from HSV‐1) leads to a decrease in the size of synapses, but an increase in synapse number (Mizoguchi et al., 2002). This observation suggests that nectin trans‐interactions are important for synaptic plasticity, and hence may be involved in learning and memory formation. Recently, nectin trans‐interactions were demonstrated to reduce endocytosis of E‐cadherin molecules, which were not involved in trans‐interactions. The process requires the binding of the small GTPase Rap1 to afadin (Hoshino et al., 2005). Nectin‐4 is upregulated in breast cancers, and soluble nectin‐4—produced by extracellular enzymatic cleavage mediated by the disintegrin ADAM17—can be detected in sera from patients with breast cancer (Fabre‐Lafay et al., 2005). For reviews of nectins see Blaschuk and Rowlands (2002), Takai and Nakanishi (2003), and Takai et al. (2003a, b).

8

Nectin‐Like Proteins

8.1 Introduction The nectin‐like proteins, necls, comprise a family of five proteins (> Table 2-19). The first member of the family to be cloned was the human poliovirus receptor (PVR), necl‐5. As the name implies, the protein is a receptor for poliovirus, and the amino acids responsible for the virus binding are highly

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. Figure 2-6 Organization of nectins in synapses. Suggested organization of nectins‐1 and ‐3, N‐cadherin, and their respective cytosolic adapter proteins at puncta adherentia junctions in synapses. (b) shows an enlarged portion of the connections presented in (a). PSD, postsynaptic density (Modified from Takai et al., 2003a)

. Table 2-19 Nomenclature for necl family CAM name Necl‐1 Necl‐3 Necl‐4 Necl‐5 SynCAM

Protein and gene synonyms BIgR, FLJ10698, IGSF4B, TSLL1 A830029E02Rik, IGSF4D IGSF4C, TSLL2 CD155, HVED, mE4, PVR, PVS, Taa1, Tage4 2900073G06Rik, 3100001I08Rik, A175N, BL2, HGNC:12378, IGSF4, IGSF4A, NECL‐2, RA175A, RA175B, RA175C, SgIGSF, ST17, TSLC1

conserved in functional PVRs of primates (Koike et al., 1992). Subsequently, a number of related proteins have been cloned. Four of these proteins, nectins‐1 to ‐4, have been placed in one family (see > Sect. 9), whereas PVR and four other proteins have been proposed to constitute another family. However, the proteins from the two families exhibit the same general structure, and proteins of both families mediate a number of Ca2þ/Mg2þ‐independent interactions. Therefore, the distinction between the two families is based solely on the ability of the proteins to bind to the cytosolic protein afadin. Thus, nectins binds to afadin, whereas nectin‐like proteins supposedly do not (> Figure 2-5) (Ikeda et al., 2003; Takai et al., 2003).

8.2 Genes Human necl‐5 was cloned in 1989 (Mendelsohn et al., 1989; Koike et al., 1990); rat and mouse necl‐5 was cloned during 1996–1997 (Chadeneau et al., 1996, 1997). Necl‐5 is not well conserved between species.


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Therefore, it has been questioned whether the genes coding for the human, mouse, and rat necl‐5 are orthologous. However, on investigations of genomic organization, expression pattern a.o., it has been concluded that the rodent necl‐5 (Tage4) indeed is orthologous to human necl‐5 (CD155; Baury et al., 2001; Ravens et al., 2003). The promoter of the gene encoding necl‐5 binds the transcription factors AP‐2a and NRF‐1 (Solecki et al., 1999, 2000), and the transcription factor Shh has been shown to activate the transcription factors Gli1 and Gli3, which subsequently stimulates transcription of necl‐5 (Solecki et al., 2002). Furthermore, the transcription factor AP‐1 stimulates Necl‐5 transcription following stimulation of the Ras–MAPK pathway (Hirota et al., 2005). Human and mouse SynCAM/Necl‐2 were cloned in 1999 and 2002, respectively (Gomyo et al., 1999; Kuramochi et al., 2001; Wakayama et al., 2001; Fukami et al., 2002), and its expression has been shown to be regulated by the transcription factor MITF (Ito et al., 2003). Human and mouse necl‐1 and ‐4 were cloned in 2001 and 2003, respectively (Fukuhara et al., 2001; Fukami et al., 2003), and the sequence of necl‐3 from humans and mice was reported directly to GenBank in 2002 (see also Takai et al., 2003). (See also > Table 2-20).

. Table 2-20 Gene information for necl family members Species Man

Mouse

Approved gene symbol IGSF4 IGSF4B IGSF4C IGSF4D PVR Igsf4a Igsf4b Igsf4c Igsf4d Pvr

Approved gene name Immunoglobulin superfamily, member 4 Immunoglobulin superfamily, member 4B Immunoglobulin superfamily, member 4C Immunoglobulin superfamily, member 4D Poliovirus receptor Immunoglobulin superfamily, member 4A Immunoglobulin superfamily, member 4B Immunoglobulin superfamily, member 4C Immunoglobulin superfamily, member 4D Poliovirus receptor

Locus 11q23.2 1q21.2–q22 19q13.32 3p12.2 19q13.2 9 B‐C 1 H3 7 A3‐B2 16 syntenic 7 A2‐B1

Database reference HGNC ID: 5951 HGNC ID: 17601 HGNC ID: 30825 HGNC ID: 29849 HGNC ID: 9705 MGI ID: 1889272 MGI ID: 2137858 MGI ID: 2449088 MGI ID: 2442722 MGI ID: 107741

8.3 Expression Necl‐1 is predominantly expressed in fetal and adult brain, the expression increasing with age. The protein is localized at cell–cell interaction sites in various cells including pyramidal cells of the hippocampus, granule cells of the cerebellum, and motor neurons of the spinal cord (Fukuhara et al., 2001; Fukami et al., 2003; Kakunaga et al., 2005; Zhou et al., 2005). Reports of the expression of SynCAM show species‐dependent variations. In rat, SynCAM is reported to be brain specific, and expressed in a developmentally regulated manner. Thus, the protein is not detectable at birth but is present in increasing amounts during the first three postnatal weeks. However, mRNA encoding SynCAM is more widely expressed than the protein itself, indicating that the expression of the protein is restricted to brain tissue because of posttranslational mechanisms (Biederer et al., 2002). In mouse, SynCAM is widely expressed during embryonic development, in the PNS and CNS and in the epithelium of various nonneuronal tissues. In the developing spinal cord the protein is expressed on axons of motor neurons and in the neural crest. The protein is also highly expressed on axons in various regions of the developing brain and sensory organs including the cortex, thalamus, habenular nucleus, olfactory system, and developing retina (Fujita et al., 2005). The protein is only expressed in the hippocampus during embryonic development (Urase et al., 2001), whereas in the telencephalon and cerebellum it exhibits an increased expression with age, with a high expression in adulthood (Ohta et al., 2005). In the adult mouse,

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SynCAM is also highly expressed in spermatogenic cells, liver, kidney, and epididymis (Wakayama et al., 2001; Shingai et al., 2003). In the brain, SynCAM colocalizes with synaptophysin but is present in both pre‐ and postsynaptic compartments (Biederer et al., 2002). Outside the nervous system the protein is located at the basolateral membrane in epithelial cells of the gall bladder and in polarized cells lining the lumen of developing mouse lung epithelium. However, the protein is not found at adherence junctions, tight junctions, or desmosomes (Fujita et al., 2003; Shingai et al., 2003). Furthermore, SynCAM in mice and humans is expressed by a subset of dendritic cells involved in interactions with T cells (Galibert et al., 2005). Necl‐4 is predominantly expressed at cell–cell interaction sites in the brain, prostate, and kidney, and also in the heart, spleen, liver, and lung (Fukuhara et al., 2001; Fukami et al., 2003; Williams et al., 2006). In the human colon cancer cell line Caco‐2 and in the kidney epithelial cell line MDCK transfected with Necl‐4, the protein is localized in the lateral parts of the plasma membrane, where it is engaged in cell–cell interactions. However, the proteins do not localize to tight junctions (Williams et al., 2006). In the developing mouse embryo necl‐5 is expressed in various regions of the CNS including the floor plate of the neural tube, the notochord (below the neural tube) as well as retinal nerve fibers, the optic nerve, and the region of the future optic chiasm (Gromeier et al., 2000). Necl‐5 is also found in lymphoid tissues, where it colocalizes with vitronectin (Lange et al., 2001), and soluble necl‐5 can be found in human serum and cerebrospinal fluid (Baury et al., 2003). Necl‐5 is reported to be overexpressed in rat colon carcinoma cell lines, tumors, and mouse intestinal adenomas (Denis, 1998).

8.4 Isoforms and Protein Structure The overall structure of necls is similar to that of nectins. Thus, the proteins all contain an extracellular domain consisting of three Ig‐modules, followed by a transmembrane segment and a short (46 amino acids) cytoplasmic tail (Fukuhara et al., 2001; Takai et al., 2003). On the basis of sequence comparison and secondary structure predictions, the N‐terminal Ig‐module is of the V‐type, the second module of the C1‐like type, and the third module of the C2 type (Du Pasquier et al., 2004). Necls‐1, ‐2, ‐3, and ‐4 have—like the nectins—a 4‐amino acid‐long motif, located at the extreme C terminus, which is capable of binding to PDZ domains (Takai et al., 2003). Necl‐5 does not contain the PDZ‐binding motif, but contains a so‐called immunoreceptor tyrosine‐based inhibitory motif, ITIM (amino acid 395–401 of human necl‐5). This motif has the consensus sequence I/V/L‐x‐Y‐x‐x‐L/V and is conserved between necl‐5 from different species (Oda et al., 2004). Necls‐1, ‐2, and ‐4 also contain a binding motif for protein 4.1, a peripheral membrane protein involved in the organization of the actin cytoskeleton (Yageta et al., 2002; Fukami et al., 2003). SynCAM exists in transmembrane and soluble isoforms, derived from alternative splicing of a single gene (Koma et al., 2004). Necl‐5 exists in four isoforms derived from alternative splicing of a single gene, the transmembrane necls‐5a‐ and ‐b (the b‐isoform being slightly shorter than the a‐isoform), and the secreted necls‐5g and ‐d (the d‐isoform being shorter than the g‐isoform) (Koike et al., 1990). The remaining necls have so far only been identified as transmembrane proteins.

8.5 Posttranslational Modifications SynCAM is known to be N‐glycosylated and contains six sites for potential N‐glycosylation. No O‐ glycosylations has been observed. Glycosylation is developmentally regulated, being most prominent during the first two postnatal weeks of mouse development (Biederer et al., 2002; Masuda et al., 2002; Fujita et al., 2003). Necl‐5 contains up to eight putative N‐glycosylation sites (Mendelsohn et al., 1989; Koike et al., 1990). Furthermore, transmembrane isoforms of necl‐5 can be serine phosphorylated (Bibb et al., 1994), and the tyrosine residue in the ITIM motif (Y398 of human necl‐5) can be phosphorylated by Src kinases (Oda et al., 2004).


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8.6 Necl–Necl Interactions Necl‐1 is reported to be involved in heterophilic trans‐interactions with SynCAM (Shingai et al., 2003). SynCAM itself forms homodimers through cis‐ (Biederer et al., 2002; Masuda et al., 2002) and trans‐ interactions (Fujita et al., 2003) (> Table 2-21). The extracellular interactions of necl‐3 have not been investigated. However, necl‐4 has been shown to form both homophilic cis‐ and trans‐interactions (Fukami et al., 2003; Williams et al., 2006).

. Table 2-21 Binding partners for necl family members Necl Necl‐1

SynCAM

Necl‐4 Necl‐5

Necl‐1 SynCAM

Necl‐5

Extracellular binding partner Necl‐1 (trans) SynCAM (trans) Nectin‐1 Nectin‐3 SynCAM (cis, trans) Necl‐1 (trans) Nectin‐3 CRTAM Necl‐4 (cis, trans) Various viruses CD44 CD96 DNAM‐1 Nectin‐3 Vitronectin 4.1N MPP3 CASK Syntenin DAL‐1 Pals2 SHP‐2 Tctex‐1

Reference Kakunaga et al. (2005) Shingai et al. (2003); Kakunaga et al. (2005) Kakunaga et al. (2005) Kakunaga et al. (2005) Biederer et al. (2002); Masuda et al. (2002); Fujita et al. (2003) Shingai et al. (2003) Ito et al. (2003) Arase et al. (2005); Galibert et al. (2005) Fukami et al. (2003); Williams et al. (2006) Koike et al. (1991); Selinka et al. (1991) Freistadt and Eberle (1997) Fuchs et al. (2004) Bottino et al. (2003); Tahara‐Hanaoka et al. (2004); Pende et al. (2005) Ikeda et al. (2003); Mueller and Wimmer (2003) Lange et al. (2001) Zhou et al. (2005) Fukuhara et al. (2003b) Biederer et al. (2002) Biederer et al. (2002) Yageta et al. (2002) Shingai et al. (2003) Oda et al. (2004) Mueller et al. (2001)

8.7 Binding Partners 8.7.1 Extracellular Binding Partners It has been demonstrated that SynCAM does not bind to L1, NCAM, or TAG‐1 (Biederer et al., 2002). However, SynCAM interacts in trans with necl‐1 and nectin‐3 (Ito et al., 2003; Shingai et al., 2003). Furthermore, SynCAM expressed by dendritic cells interacts with CRTAM, an IgSF receptor expressed on a subset of natural killer cells and T cells. The interactions between CRTAM and SynCAM have higher affinity than the homophilic SynCAM interactions, and CRTAM disrupts homophilic SynCAM and heterophilic SynCAM/Necl‐1 interactions, indicating that the binding sites on SynCAM for CRTAM, Necl‐1, and SynCAM are overlapping (Arase et al., 2005; Boles et al., 2005; Galibert et al., 2005).

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Necl‐5 is originally known for its ability to bind various viruses including poliovirus. The viruses bind to the first Ig‐module of necl‐5 (Koike et al., 1991; Selinka et al., 1991) in a manner independent of the glycosylation of the domain (Koike et al., 1992; Zibert and Wimmer, 1992). In addition, necl‐5 interacts in trans with the hyaluronic acid receptor CD44, the leukocyte adhesion molecules CD96 and DNAM‐1, and nectin‐3 (Freistadt and Eberle, 1997; Bottino et al., 2003; Ikeda et al., 2003; Mueller and Wimmer, 2003; Fuchs et al., 2004; Tahara‐Hanaoka et al., 2004; Pende et al., 2005), and necl‐5 binds to the ECM molecule vitronectin with a Kd around 72 nM (Lange et al., 2001) (> Table 2-21).

8.7.2 Intracellular Binding Partners Necl‐1 binds the protein 4.1N, a member of the protein 4.1 family (Zhou et al., 2005). The PDZ‐interacting motif of SynCAM interacts with the proteins MPP3, CASK, and syntenin (Biederer et al., 2002; Fukuhara et al., 2003b), and the protein also interacts with DAL‐1 (another protein belonging to the protein 4.1 family) (Yageta et al., 2002), and with Pals2, a membrane‐associated guanylate kinase (Shingai et al., 2003). The cytoplasmic tail of necl‐5 binds to SHP‐2 and Tctex‐1. SHP‐2 is a tyrosine phosphatase that becomes activated by the interaction with necl‐5 (Oda et al., 2004). Tctex‐1 is a light chain of the dynein motor complex that facilitates retrograde transport of endocytic vesicles and membranous organelles (Mueller et al., 2001). Necl‐5 has also been reported to colocalize and associate with integrin avb5. However, the nature of the interaction, i.e., extracellular or intracellular, direct or indirect, has not been shown (Ikeda et al., 2004) (> Table 2-21).

8.8 Signaling Binding of anti‐necl‐5 antibodies, poliovirus, or DNAM‐1 to necl‐5 induces phosphorylation of the tyrosine residue located in the ITIM motif in the cytoplasmic tail of this protein. The phosphorylation is facilitated by Src kinases, and the phosphorylated tyrosine residue recruits the tyrosine phosphatase SHP‐2, which becomes activated upon the interaction. Subsequently, focal adhesion kinase, FAK, and paxillin become dephosphorylated. The dephosphorylation of FAK leads to a reduction in its kinase activity, and this is possibly the reason for the reduction in focal adhesion‐mediated cell attachment observed by stimulation of necl‐5 (Oda et al., 2004). Necl‐5 also activates the small GTPases Cdc42 and Rac, which induce the formation of filopodia and lamellipodia, respectively (Ikeda et al., 2004). Necl‐5 also modulates PDGF‐stimulated cell proliferation. Necl‐5 acts downstream of the PDGF‐ receptor but upstream of Ras. Expression of recombinant necl‐5 lacking a part of the cytoplasmic tail reduces PDGF‐mediated cell proliferation by reducing activation of Ras–Raf–MEK–ERK signaling, whereas necl‐5 enhances proliferation by activating the ERK kinases and by upregulating cyclins D2 and E and downregulating p27Kip1 (Kakunaga et al., 2004).

8.9 Functions All necls seem to mediate cell–cell and/or cell–ECM interactions. Cells overexpressing SynCAM demonstrate increased aggregation and adhesion (Masuda et al., 2002; Ito et al., 2003). Secreted isoforms of SynCAM bind to transmembrane isoforms of SynCAM and inhibit cell–cell interactions in a dose‐ dependent manner (Koma et al., 2004). In coculture systems, expression of full‐length SynCAM in the confluent layer of nonneuronal ‘‘feeder’’ cells induces synapse formation in hippocampal neurons growing on top of the ‘‘feeder’’ layer, whereas expression of the cytoplasmic tail of SynCAM in neurons inhibits synapse formation (Biederer et al., 2002). SynCAM and necls‐1 and ‐4 are believed to be tumor suppressors. Thus, loss of expression of necl‐1 or ‐4 has been observed in prostate cancer and in various gliomas (Fukuhara et al., 2001), and in some cancer


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cells SynCAM is expressed in a truncated version lacking a part of the cytoplasmic tail, indicating that this part of the molecule is important for its tumor‐suppressive function (Kuramochi et al., 2001). Furthermore, SynCAM is an inhibitor of metastasis, possibly because of its ability to mediate cell–cell interactions (Yageta et al., 2002). Alterations in SynCAM expression in various cancer cells have been shown to correlate with the methylation of the SynCAM promoter (Kuramochi et al., 2001). Furthermore, the interaction between SynCAM and CRTAM modulates cytotoxicity, migration, interferon g secretion, and interleukin‐22 expression of NK cells and of CD8þ T cells (Arase et al., 2005; Boles et al., 2005; Galibert et al., 2005). Finally, the expression pattern of SynCAM in the nervous system indicates that it may be involved in the migration, differentiation, and fasciculation of neurons (Fujita et al., 2005). Necl‐5 is also important for immune responses, where trans‐interactions between necl‐5 and DNAM‐1 or CD96 stimulates the cytotoxic response and cytokine secretion of NK cells and T cells (Bottino et al., 2003; Fuchs et al., 2004; Tahara‐Hanaoka et al., 2004). The interaction between necl‐5 and DNAM‐1 is also important for the regulation of the transendothelial migration of monocytes (Reymond et al., 2004). Necl‐5 is also a modulator of cell migration. The protein colocalizes with integrin avb3 at the leading edge of migrating fibroblasts, and expression of necl‐5 reduces cell adhesion (focal adhesions), but induces the formation of filopodia and lamellipodia and enhances serum‐ and PDGF‐induced proliferation and motility in coculture systems and trans‐well assays with fibronectin‐coated membranes. The extracellular part of necl‐5 is only necessary for modulation of directional cell migration, whereas removal of the cytoplasmic tail affects both directional and random cell motility (Ikeda et al., 2004; Kakunaga et al., 2004; Oda et al., 2004). For recent reviews of necls see Abbas (2003), Campadelli‐Fiume et al. (2000), Takai et al. (2003), Watabe et al. (2003), and Yamagata et al. (2003).

9

The IgLON Family

9.1 Introduction OBCAM (opioid‐binding cell adhesion molecule) was identified as an opioid receptor in 1986 (Cho et al., 1986). However, human OBCAM was not cloned until 1995 (Shark and Lee, 1995). The related proteins LAMP (limbic system‐associated membrane protein) and neurotrimin were cloned the same year (Pimenta et al., 1995; Struyk et al., 1995), and the three proteins were, on basis of their strong similarities, placed in a separate family named IgLON (IgSF containing LAMP, OBCAM, and neurotrimin). More recently, an additional member of the family, Kilon (kindred of IgLON), was cloned (Funatsu et al., 1999) (> Table 2-22). . Table 2-22 Nomenclature for IgLON family CAM name LAMP Neurotractin Neurotrimin OBCAM

Protein and gene synonyms B130007O04Rik, Lam 5330422G01Rik, KILON, MGC46680, NEGR1, Ntra 6230410L23Rik MGC:99974,OPCM

9.2 Genes The mouse gene that encodes LAMP has been investigated in detail. It is composed of 11 exons. The gene is unusual in having two different exon 1, which are separated by 1.6 megabase pairs (Mbp) and contain separate promoter regions. In addition, alternative splicing of two small exons results in isoforms containing or lacking a 23‐amino acid‐long extracellular membrane‐proximal insert (Pimenta et al., 1998;

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. Table 2-23 Gene information for IgLON family members Species Man

Approved gene symbol HNT LSAMP NEGR1 OPCML

Mouse

Hnt Lsamp Negr1 Opcml

Approved gene name Neurotrimin Limbic system‐associated membrane protein Neuronal growth regulator 1 Opioid‐binding protein/cell adhesion molecule‐like Neurotrimin Limbic system‐associated membrane protein Neuronal growth regulator 1 Opioid binding protein/cell adhesion molecule‐like

Locus 11q25 3q13.2–q21

Database reference Locus ID: 50863 HGNC ID: 6705

1p31.1 11q24.3– 11q24.3 9 syntenic 16 B5

HGNC ID: 17302 HGNC ID: 8143

3 syntenic 9 10.0

MGI: 2444846 MGI: 97397

MGI: 2446259 MGI: 1261760

Pimenta, and Levitt, 2004). In the human brain, two LAMP mRNA transcripts 1.6 and 8.0 kb have been identified (Pimenta et al., 1996). (See also > Table 2-23).

9.3 Expression OBCAM is predominantly expressed in the brain, but also found in other tissues including the spleen. In rats, the protein is detected in the brain around E16. Postnatally, the expression of the protein increases and is sustained at a high level during adulthood. OBCAM is expressed in gray matter but is absent in white matter, and during development found in the olfactory bulb, cerebral cortex, hippocampus, striatum, septum, thalamus, cerebellar cortex, cerebellar nuclei, nuclei in the brain stem, and in the spinal cord (Hachisuka et al., 1999, 2000). In the adult rat, OBCAM is predominantly found in the cerebral cortex and hippocampus, and to a lesser extent in the olfactory bulb and diencephalon, whereas the expression in the cerebellum, medulla oblongata, and spinal cord is low (Miyata et al., 2003a). The protein is generally expressed as a 51‐kDa isoform, but in the olfactory bulb and cerebellum it is predominantly expressed as a 46‐kDa isoform (Miyata et al., 2003a). In fully differentiated cortical and hippocampal neurons, OBCAM is expressed on the somata and dendrites, but not on the axonal fibers (Miyata et al., 2000, 2003b, c). LAMP is expressed by neurons of the limbic system (Cote et al., 1995). In the rat brain, LAMP is detectable around E15, and early in fetal development the proteins are expressed on growing axons (Horton and Levitt, 1988). In the adult rat, LAMP is exclusively located postsynaptically on somata and dendrites and is not found on axons. In hippocampal cultures, LAMP is expressed by neurofilament‐ and MAP2‐ expressing neurites, but not by GFAP‐expressing astroglial cells (Zacco et al., 1990). The expression of neurotrimin during rat development is restricted to the nervous system including the spinal cord, dorsal root ganglia, and forebrain. At P0 the protein is expressed in the olfactory bulb, pyriform cortex, midbrain, thalamus, hypothalamus, hippocampus, basal ganglia, and cortical plate. At this time point it is found in the CA1, CA2, and CA3 hippocampal regions, but later, at P14, it is only expressed in the CA1 region (Struyk et al., 1995). During development neurotrimin is located on unmyelinated axons, but later it is predominantly located at excitatory synaptic contact sites (Chen et al., 2001). The expression of Kilon is also restricted to the nervous system. During rat development, the proteins are detectable around E16, after which the level gradually increases. Kilon is expressed in the cerebral cortex, brain stem and hippocampus and—in lower amounts—in the cerebellum. In the adult rat, Kilon is


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predominantly expressed in the olfactory bulb, cerebral cortex, diencephalon, hippocampus, and cerebellum (Funatsu et al., 1999; Miyata et al., 2003a). Kilon is predominantly located postsynaptically on the somata and dendrites of neurons (Funatsu et al., 1999; Miyata et al., 2003a, c).

9.4 Isoforms and Protein Structure All IgLON proteins are glycosylphosphatidylinositol (GPI)‐linked receptors composed of three Ig‐modules of the C2 type (Schofield et al., 1989; Wick et al., 1996; Funatsu et al., 1999). The proteins are highly conserved with sizes ranging from 338 to 348 amino acids. Human and rat OBCAM and LAMP are 98% and 94% identical, respectively (Shark and Lee, 1995; Pimenta et al., 1996), and in rat, Kilon is 56%, 49%, and 48% similar to LAMP, OBCAM, and neurotrimin, respectively, whereas LAMP is 56%, and 55% similar to OBCAM and neurotrimin, respectively, and OBCAM is 77% similar to neurotrimin (Pimenta et al., 1996; Funatsu et al., 1999). The first Ig‐modules of OBCAM and neurotrimin are 94% identical, and show strong homology to the first Ig‐module of FGFR, whereas the second and third Ig‐modules of these proteins show strong homology to IgIV and IgIII of NCAM, respectively (Struyk et al., 1995).

9.5 Posttranslational Modifications All IgLON proteins have 6–8 sites for potential N‐linked glycosylation (Struyk et al., 1995; Hachisuka et al., 1996; Pimenta et al., 1996; Funatsu et al., 1999). Western blotting has revealed OBCAM as two bands 51 and 58 kDa, respectively, the difference in size being a result of glycosylation (Hachisuka et al., 1996). The glycosylation of LAMP does not include the HNK‐1 epitope, but does contain high mannose sugars and a minor amount of sialic acid (Zacco et al., 1990).

9.6 Interactions Within the IgLON Family It has been reported that neurotrimin can form cis‐homodimers and multimers, and that both LAMP and neurotrimin mediate homophilic trans‐interactions, and also can interact with each other in trans‐heterophilic interactions (Zhukareva and Levitt, 1995; Gil et al., 1998, 2002). Kilon is reported to interact homophilically and heterophilically with OBCAM. Furthermore, OBCAM is involved in homophilic trans‐interactions and heterophilic trans‐interactions with CEPU‐1 (chicken neurotrimin) (McNamee et al., 2002; Miyata et al., 2003a). Recently, it was proposed that the individual members of the IgLON family are subunits of cis‐homo‐ or heterodimers in the plasma membrane (named Diglons), and the previously reported trans‐interactions are believed to occur between Diglons rather than between individual IgLONs (> Table 2-24) (Reed et al., 2004). In LAMP, the homophilic interaction is believed to be mediated by the first Ig‐module (Eagleson et al., 2003). . Table 2-24 Binding partners for IgLON family members CAM IgLONs

Extracellular binding partner IgLONs (cis, trans)

OBCAM

Opioid alkaloids

Reference Zhukareva and Levitt (1995); Gil et al. (1998, 2002); McNamee et al. (2002); Eagleson et al. (2003); Miyata et al. (2003a); Reed et al. (2004) Ann et al. (1992); Govitrapong et al. (1993); Shark and Lee (1995)

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9.7 Heterophilic Binding Partners OBCAM was originally identified as a receptor for opioid alkaloids in the presence of acidic lipids (> Table 2-24) (Shark and Lee, 1995).

9.8 Signaling The expression of OBCAM has been demonstrated to be important for opioid signaling mediated via heterotrimeric G proteins, and a reduction in cellular OBCAM expression leads to a reduction in the binding of opioids to cells (Ann et al., 1992; Govitrapong et al., 1993). LAMP can induce neurite outgrowth in a manner involving L‐type calcium channels, and incubation of cells in the presence of soluble LAMP can induce a sustained increase in intracellular Ca2þ (Zhukareva et al., 1997).

9.9 Function IgLONs seem in part to function as axonal guidance molecules. Thus, LAMP appears on limbic system axons in a manner correlating with the formation of pathways created by these axons (Horton and Levitt, 1988), and during development OBCAM is highly expressed on the surface of growing axons (Hachisuka et al., 2000). The molecules can also act as repulsive axonal guidance molecules. Thus, OBCAM–neurotrimin heterodimers mediate trans‐inhibition of neurite outgrowth from cerebellar granule cells, and LAMP inhibits neurite outgrowth of (nonlimbic) neurotrimin‐expressing dorsal root ganglion neurons, whereas limbic neurons aggregate and extend neurites, when plated on a substrate coated with LAMP (Zhukareva and Levitt, 1995; Mann et al., 1998; Gil et al., 2002; Reed et al., 2004). Furthermore, neurotrimin may be important for both axonal fasciculation and formation and stabilization of excitatory synapses (Chen et al., 2001).

10

Activated Leukocyte‐Cell Adhesion Molecule

10.1 Introduction ALCAM (activated leukocyte‐cell adhesion molecule) was originally cloned in chicken, where it is known by various names e.g., BEN and DM‐GRASP. The mouse orthologue was cloned in 1994 and was shown to be 76% identical to chicken ALCAM. Later, the human and rat orthologues were cloned (Kanki et al., 1994; Bowen et al., 1995; Uchida et al., 1997; Stephan et al., 1999) (> Table 2-25). . Table 2-25 Nomenclature for ALCAM CAM name ALCAM

Protein and gene synonyms AG 2117, BEN, CD166, DM‐GRASP, HCA, hematopoietic cell antigen, MEMD, MGC:27910, MuSC, SC1

Human ALCAM exhibits homology with the zebrafish orthologue of ALCAM, neurolin (38% identity, 55% similarity), with the receptor RAGE (28% identity, 43% similarity with human RAGE), MCAM (23% identity, 49% similarity with human MCAM) (Bowen et al., 1995) (see > Sect. 13), and the laminin‐ interacting adhesion molecule B‐CAM (26% amino acid identity) (Campbell et al., 1994).


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10.2 Gene In activated leukocytes, ALCAM mRNAs 5.2, 8.5, and 10 kb have been reported (Bowen et al., 1995), whereas a 1.8 kb mRNA has been reported in testis. However, in most tissues, the expression is seen as a single mRNA of 5 kb (Uchida et al., 1997). Human ALCAM seems to exist in at least two isoforms formed by alternative splicing. The two isoforms are identical except for a deletion between amino acids 503 and 516 (immediately before the transmembrane segment) in one of the isoforms (Uchida et al., 1997). (See also > Table 2-26).

. Table 2-26 Gene information for ALCAM Species Man

Approved gene symbol ALCAM

Mouse

Alcam

Approved gene name Activated leukocyte‐cell adhesion molecule

Locus 3q13.1


16 B5

MGI ID: 1313266

10.3 Expression ALCAM originally got its name owing to its expression in activated leukocytes (Bowen et al., 1995). However, later it was demonstrated also to be expressed in many other tissues. In humans, ALCAM is expressed in the blastocysts and the endometrium, suggesting that ALCAM‐ mediated adhesion is important for the initial attachment of the embryo to the maternal endometrium (Fujiwara et al., 2003). In the 7‐week‐old human embryo, ALCAM is expressed on cell bodies and axons of dorsal root ganglion neurons and motor neurons in the neural tube, but it is also expressed in nonneuronal tissues including the notochord, heart, lung, kidneys, and 30% of hematopoietic cells. In humans, ALCAM is predominantly expressed in the ovaries and the prostate in adulthood, but it is also expressed in all regions of the brain as well as other nonneuronal tissues (Karagogeos et al., 1997; Uchida et al., 1997; Stephan et al., 1999). In the rat at E10, ALCAM has been demonstrated to be expressed in the notochord. Later, the protein is also expressed in the floor plate, heart, lungs, and skin, and around E15–E18 the protein is ubiquitously expressed. In the adult rat, the protein is found predominantly in the brain, kidneys, and pancreas (Stephan et al., 1999). During development in mice, ALCAM is expressed in the epithelia of both ectodermal and endodermal origins and in various neuronal tissues including the spinal cord, dorsal root ganglia, and trigeminal ganglia. Furthermore, the protein appears to be important for facial development, where among other tissues it is highly expressed in the developing teeth (Fraboulet et al., 2000, 2003). In a study of the expression of the protein in cows it was shown to be constitutively expressed in the autonomous nervous systems (Konno et al., 2001). ALCAM has been proposed to be a marker for several types of cancers. Thus, the expression of the protein has been shown to correlate with the aggregation and metastatic capacity of human melanoma cells (van Kempen et al., 2000), and in a transcription analysis of prostate cancer specimens, the expression of ALCAM was shown to be upregulated in 22% of cases (demonstrating a 2‐ to 3.8‐fold increase). Immunohistochemically, the protein displayed an increased expression in 81% of examined specimens (Kristiansen et al., 2003a). In an in vitro study of seven prostate cancer cell lines, it was demonstrated that ALCAM only is localized to the membrane if the cells express a‐catenin. Absence of a‐catenin and, to a lesser degree, E‐cadherin leads to a cytoplasmic localization of ALCAM (Tomita et al., 2000). Finally, the expression of ALCAM is increased in colorectal and esophageal carcinomas (Weichert et al., 2004; Verma et al., 2005).

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10.4 Isoforms and Protein Structure The two isoforms of human ALCAM are 584 and 571 amino acid long, respectively. The proteins consist of a 27‐amino acid‐long signal peptide, a 500‐amino acid‐long extracellular domain, a 24‐amino acid‐long transmembrane region, and a 32‐amino acid‐long cytoplasmic tail. The extracellular domain consists of five Ig‐modules of which the two most membrane‐distal modules are of the V‐type, and the remaining three modules of the C2‐type (Bowen et al., 1995; Uchida et al., 1997).

10.5 Posttranslational Modifications ALCAM contains eight—ten potential sites for N‐linked glycosylation (Bowen et al., 1995; Uchida et al., 1997). Glycosylation on ALCAM includes the a2,8‐linked disialic acid (diSia) epitope, a type of glycosylation, which probably is mediated by the enzyme a2,8‐sialyltransferase III (Sato et al., 2002).

10.6 Homophilic Interactions ALCAM is involved in homophilic trans‐interactions with a Kd around 29–48 mM (Uchida et al., 1997; Hassan et al., 2004). The binding is mediated by a reciprocal interaction between the first Ig‐modules of two ALCAM molecules. However, the avidity of the interaction seems to be dependent on the two most membrane‐proximal Ig‐modules (number four and five), and therefore a model has been proposed where ALCAM interacts homophilically in cis with the fourth and fifth Ig‐modules and in trans with the first Ig‐module (van Kempen et al., 2001) (> Table 2-27). . Table 2-27 Binding partners for ALCAM CAM ALCAM

Extracellular binding partner ALCAM (cis) ALCAM (trans) CD6 (trans) NrCAM

Reference van Kempen et al. (2001) Uchida et al. (1997); Nelissen et al. (2000); Hassan et al. (2004); Zimmerman et al. (2004) Bajorath et al. (1995); Whitney et al. (1995); Bowen et al. (1995, 1996, 2000); Skonier et al. (1996a, b, 1997); Bodian et al. (1997), Hassan et al. (2004) DeBernado and Chang (1996)

Homophilic ALCAM trans‐interactions are reported to be affected by alterations in the organization and dynamics of the actin cytoskeleton in a manner dependent on the activity of PKCa (Nelissen et al., 2000; Zimmerman et al., 2004).

10.7 Heterophilic Binding Partners 10.7.1 Extracellular Binding Partners ALCAM interacts heterophilically in trans with CD6 with a Kd around 0.4–1.0 mM (Bowen et al., 1995; Hassan et al., 2004) (> Table 2-27). CD6 is a member of the scavenger receptor family, predominantly expressed by thymocytes and certain types of B‐cells and T‐cells. Extracellularly, CD6 is composed of three scavenger receptor cysteine‐rich (SRCR) modules. ALCAM and CD6 interact in a 1:1 ratio, and the binding takes place between the first Ig‐module of ALCAM and the third (most membrane‐proximal) SRCR module of CD6 (Bajorath et al., 1995;


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Whitney et al., 1995; Bowen et al., 1996). Residues responsible for the interaction include F26, K28, F40, F43, K48, Y53, D54, M87, T90, and E91 of ALCAM and Q352, N345, N346, and N348 of CD6 (Skonier et al., 1996a, b, 1997; Bodian et al., 1997) (see reviews by Aruffo et al. (1997) and Bowen et al. (2000)). In the chick, ALCAM has been shown to interact in trans or cis with NrCAM (DeBernado and Chang, 1996) (see also > Sect. 19).

10.8 Function In vitro experiments have demonstrated that incubation of embryonic pancreatic ductal epithelial cells with a recombinant construct of soluble ALCAM leads to a dose‐dependent inhibition of cell growth and a change in the degree of tyrosine phosphorylation of cytosolic proteins (Stephan et al., 1999). Furthermore, ALCAM has been related to the process of neurite outgrowth. Thus, induction of cellular differentiation in N2a neuroblastoma cells by retinoic acid leads to an increased expression by ALCAM with the diSia glycoepitope, and simultaneous treatment with antibodies against this epitope impairs neurite outgrowth (Sato et al., 2002). ALCAM‐null mice are viable, fertile, and do not display any external morphological defects. However, they do demonstrate defects in the axon guidance and fasciculation of certain groups of neurons, and they also have morphological alterations in the retina (Weiner et al., 2004). Recently, ALCAM was demonstrated to undergo clathrin‐mediated endocytosis. However, endocytosed ALCAM is not degraded in lysosomes, but recycled to the plasma membrane (Piazza et al., 2005). For a recent review of ALCAM see Swart et al. (2005).

11

Melanoma Cell Adhesion Molecule

11.1 Introduction MCAM (melanoma cell adhesion molecule) is related to ALCAM (23% identity between human forms) (Bowen et al., 1995) (see > Sect. 12) and B‐CAM (an adhesion molecule known to interact with laminin; 31% amino acid identity between human forms) (Campbell et al., 1994). These three proteins all contain an extracellular domain composed of two V‐type Ig‐modules followed by three C2‐type Ig‐modules. MCAM, an orthologue of the chicken protein gicerin (cloned by Taira and coworkers (1994)), is also known by this as well as several other names (Shih et al., 1994; Schon et al., 2005) (see > Table 2-28). Human, mouse, and rat MCAMs were cloned in 1989, 2001, and 2004, respectively (Lehmann et al., 1989; Yang et al., 2001b; Taira et al., 2004). . Table 2-28 Nomenclature for MCAM CAM name MCAM

Protein and gene synonyms CD146, gicerin, HEMCAM, melanoma‐associated antigen A32, Mel‐CAM, MUC18, S‐endo 1 endothelial‐associated antigen

11.2 Genes The human and mouse MCAM genes (> Table 2-29) both contain 16 exons, and results in mRNAs 3.3 and 3.0 kb, respectively (Sers et al., 1993; Yang et al., 2001b). The human MCAM promoter lacks conventional TATA and CAAT boxes, but contains four SP‐1 elements, two AP‐2 elements, one

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. Table 2-29 Gene information for MCAM Species Man Mouse

Approved gene symbol MCAM Mcam

Approved gene name Melanoma cell adhesion molecule

Locus 11q23.3 9 syntenic

Database reference HGNC: 6934 MGI: 1933966

CRE, a c‐Myb element as well as CarG‐box motifs. The CRE and an AP‐2 element located at 131 and 302 bp have a stimulatory effect on expression, whereas an AP‐2 site at 23 bp has an inhibitory effect (Sers et al., 1993; Mintz‐Weber and Johnson, 2000). The murine Mcam promoter also contains a CRE site, and a cellular increase in cAMP leads to a stimulation of MCAM expression (Kohama et al., 2005).

11.3 Expression In humans, MCAM is expressed in smooth muscles and abundantly in endothelial cells of various origins, where it is a component of endothelial junctions (Shih et al., 1994, 1997; Bardin et al., 2001). Within the adult human CNS, post mortem studies of human brain tissue indicate that MCAM mainly is expressed on the vasculature within the CNS (Schwarz et al., 1998). MCAM was originally identified as an integral membrane glycoprotein in human melanomas, and the protein serves as a prognostic marker for primary melanomas (Pacifico et al., 2005). The protein is expressed in more than 70% of metastatic melanomas, but not in normal melanocytes, and the invasiveness of melanomas correlates with the cellular amount of MCAM (Aldrian et al., 2003). It has also been found in the majority of investigated mucoepidermoid carcinomas (MEC) (Pires et al., 2003), and it is expressed in many nonsmall cell lung cancers (NSCLC) (Kristiansen et al., 2003b). Furthermore, it is highly upregulated in primary bronchial epithelial cells derived from patients with chronic obstructive pulmonary disease (COPD), a disease induced by tobacco abuse (Schulz et al., 2003). In contrast, the protein is not expressed by squamous cell carcinomas (SCC) (Pires et al., 2003) and it is downregulated in hemangiomas, the most common tumour of endothelial origin (Li et al., 2003a). In rat, MCAM is highly expressed in lungs, heart, and smooth muscles. However, the protein is also expressed in all brain regions. The expression is strongest during development, but still detectable in the adult CNS, and in contrast to the general level of expression in the brain, the expression of MCAM increases in the hippocampus after birth. In the hippocampus of the adult rat, MCAM is predominantly expressed in the inner molecular layer just outside of the granule cells of the dentate gyrus (Taira et al., 2004). The protein is also expressed by the rat pheochromocytoma cell line PC12 (Taira et al., 2005). In a melanoma cell line transfected with MCAM, the protein has been demonstrated to localize to microvilli (Okumura et al., 2001). The chicken orthologue of MCAM, gicerin, is highly expressed in the embryonic kidney, but also found in the developing nervous system including the retina, optic tectum, cerebellum, spinal cord, sciatic nerves, and the dorsal root ganglion (Taira et al., 1995, 2004; Hiroi et al., 2003). The expression of the protein declines after hatching (Tsukamoto et al., 1997; Hiroi et al., 2003).

11.4 Isoforms and Protein Structure Human and murine MCAMs consist of up to 646 and 648 amino acids, respectively (Yang et al., 2001b). The proteins are composed of five extracellular Ig‐modules (V‐V‐C2‐C2‐C2) followed by a transmembrane region and a short cytoplasmic tail. Alternative splicing of exon 15 results in two isoforms, l‐ and s‐MCAM (‘‘large’’ and ‘‘small’’), containing a cytoplasmic tail of 63 and 21 amino acids, respectively (Lehmann et al., 1989; Taira et al., 1995; Kohama et al., 2005). Furthermore, cultured human endothelial cells have been demonstrated to shed a soluble isoform of MCAM to the culture media. This isoform is 10 kDa smaller than transmembrane MCAM (Bardin et al., 1998).


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11.5 Posttranslational Modifications MCAM has eight potential N‐glycosylation sites (Lehmann et al., 1989), and the protein contains carbohydrates containing sialic acid and the HNK‐1 epitope (a carbohydrate epitope expressed on a number of CAMs, including NCAM, L1, TAG‐1, and P0) (Shih et al., 1994).

11.6 Homophilic Interactions It is still not clear whether MCAM is able to form homophilic trans‐interactions (see below).

11.7 Heterophilic Binding Partners 11.7.1 Extracellular Binding Partners Expression of MCAM in MCAM‐negative melanoma cell lines results in cellular aggregation. However, MCAM‐negative melanoma cells can adhere to immobilized MCAM. Furthermore, MCAM‐negative melanomas are able to form heterotypic cell aggregates when mixed with MCAM‐positive but not MCAM‐negative endothelial cells. Expression of MCAM in an MCAM‐negative colorectal cell line did not lead to increased cell–cell interactions. Together, these observations imply that MCAM may not form homophilic trans‐interactions, but instead mediates cell–cell interactions through trans‐heterophilic protein interactions with an unidentified counterreceptor. Furthermore, they imply that this counterreceptor often is expressed on melanoma cells, but not on endothelial cells (Johnson et al., 1997; Shih et al., 1997). Although MCAM contains a potential glycosaminoglycan‐binding site the protein does not appear to bind glycosaminoglycans (Johnson et al., 1997; Shih et al., 1997). The chicken orthologue of MCAM, gicerin, binds the laminin‐like matrix protein NOF (Taira et al., 1994, 1995).

11.7.2 Intracellular Binding Partners MCAM has been reported to bind the Src family kinase Fyn (Anfosso et al., 1998) and l‐MCAM but not s‐MCAM, which can bind moesin, a protein enriched in the microvilli (Okumura et al., 2001) (> Table 2-30).

. Table 2-30 Binding partners for MCAM CAM MCAM

Intracellular binding partner Fyn Moesin

Reference Anfosso et al. (1998) Okumura et al. (2001)

11.8 Function Rat E18 hippocampal neurons cultured on a recombinant, chimeric MCAM‐Fc substrate produce longer neurites than cells cultured on polylysine, but not as long as the neurites produced when cells are grown on laminin (Taira et al., 2004). Likewise, rat PC12 cells adhere to a recombinant, chimeric Fc‐MCAM substrate, and these trans‐MCAM interactions promote NGF‐induced neurite outgrowth. Furthermore, NGF treatment induces MCAM expression in PC12 cells (Taira et al., 2005). In rat hippocampus, expression of MCAM (predominantly l‐MCAM) has been shown to increase in response to maximal electroconvulsive seizures (MECS) (Taira et al., 2004), and injury of neurons in the hypoglossal nucleus also leads to an increase in MCAM expression, predominantly in reactive astrocytes

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surrounding the injured neurons. These results suggest a role for MCAM in relation to neural regeneration (Li et al., 1999). Human serum contains soluble forms of MCAM. In patients with chronic renal failure (CRF), the amount of soluble MCAM and the expression of MCAM in endothelial cells in the kidneys increase significantly, and it has been suggested that the upregulation of MCAM is a protective mechanism against endothelial loss in relation to renal dysfunction (Bardin et al., 2003; Daniel et al., 2005; Malyszko et al., 2005). MCAM generally promotes tumour growth and metastasis. Thus, transfection of the mouse fibroblastoid cell line L929 with MCAM stimulates cell proliferation and leads to invasion and metastasis subsequent to tissue implantation (Tsukamoto et al., 2003). Consistently, the level of MCAM in nine mouse melanoma cell lines was shown to be directly proportional to the ability of the cells to form metastatic colonies in lungs (Yang et al., 2001b). Increased MCAM expression in metastatic cells correlates with the loss of expression of the transcription factor AP‐2, and the tumorigenicity and metastatic potential of cells introduced into nude mice can be inhibited by the reexpression of AP‐2 (Jean et al., 1998). In contrast, the transcription factor CREB stimulates MCAM expression, and transfection of a human melanoma cell line with dominant negative CREB has been shown to reduce the invasiveness of these cells (Jean and Bar‐Eli, 2000). In a rat aortic smooth muscle cell line, exposure to IGF‐1, but not PDGF‐BB or bFGF, induced the expression of MCAM (Okumura et al., 2004). In human umbilical vein endothelial cells, binding of monoclonal anti‐MCAM antibodies to MCAM induces Ca2þ influx, and leads to the association of MCAM with Fyn and tyrosine phosphorylation of PLCg, Pyk2, p130Cas, FAK as well as association of FAK with paxillin (Anfosso et al., 1998, 2001). Thus, extracellular MCAM‐mediated protein interactions can lead to induction of intracellular signal transduction and cytoskeletal reorganization.

12

Myelin‐Associated Glycoprotein

12.1 Introduction The myelin‐associated glycoprotein (MAG) is a protein of relatively low abundance ( Table 2-31). . Table 2-31 Nomenclature for MAG CAM name MAG

Protein and gene synonyms Gma, SIGLEC‐4A

12.2 Genes The gene encoding MAG has been mapped to chromosome 19 in humans and to chromosome 7 in mice (> Table 2-32) (Barton et al., 1987). The promoter of the rat Mag has been studied by Laszkiewicz et al. (1997). . Table 2-32 Gene information for MAG Species Man Mouse

Approved gene symbol MAG Mag

Approved gene name Myelin‐associated glycoprotein

Locus 19q13.1 7 11.0



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12.3 Expression The expression of MAG is restricted to the PNS and CNS where its expression pattern is regulated in a spatial and temporal manner. MAG is expressed in oligodendrocytes and Schwann cells and the expression of different MAG isoforms is developmentally regulated (see below). In the PNS, NCAM and L1 are present in neurons and Schwann cells prior to myelination but are downregulated as myelination starts. At roughly the same time MAG and P0 become detectable (Martini et al., 1995). Before myelin compaction, the subcellular location of MAG and P0 is identical. After compaction however, MAG and P0 are located in a complementary manner, where MAG is confined to noncompacted areas of the Schwann cells and P0 is restricted to the compacted areas (see also > Sect. 4) (reviewed by Martini et al., 1994). In rat and mouse brains, mRNA encoding one isoform of MAG, L‐MAG, is expressed earlier during myelination and in higher amounts than another isoform, S‐MAG, whereas S‐MAG becomes the predominant expressed isoform later in development when the rate of myelination is decreased (Frail and Braun, 1984). Consistently, a more recent study has demonstrated that L‐MAG is expressed in larger amounts in oligodendrocytes before the onset of myelination than at later stages (Keita et al., 2002).

12.4 Isoforms and Protein Structure MAG consists of one Ig‐module of the V‐type followed by four Ig‐modules of the C2‐type, a single transmembrane segment, and a cytoplasmic tail. Alternative splicing of the Mag transcript generates two isoforms, which differ in the length of the cytoplasmic tail. The two isoforms, termed S‐MAG and L‐ MAG (‘‘small’’ and ‘‘large’’), respectively, have molecular weights of 57 and 72 kDa and cytoplasmic tails of 45 and 90 amino acids, respectively (Salzer et al., 1987). S‐MAG is primarily found in the adult nervous system while L‐MAG is synthesized during myelination in the developing nervous system (Pedraza et al., 1991).

12.5 Posttranslational Modifications MAG contains eight sites for N‐linked glycosylation, and both isoforms are heavily glycosylated, as much as 30% of the molecular weight being comprised by sugar residues (Brady and Quarles, 1988; Pedraza et al., 1991). MAG may be palmitoylated at a cysteine residue in the membrane‐spanning region (Pedraza et al., 1990) and contains putative phosphorylation sites in the intracellular domain (Edwards et al., 1989).

12.6 Homophilic Interactions MAG has not been reported to interact homophilically.

12.7 Heterophilic Binding Partners MAG is able to bind sialic acid, and this binding is believed to be of importance for the binding of MAG to neurons (Filbin, 1995; Tang et al., 1997; Kleene and Schachner, 2004). A definite protein receptor for MAG remains to be identified, but MAG has been demonstrated to interact with the brain gangliosides GD1a and GT1b (Yang et al., 1996), and the interaction between MAG and GT1b may be responsible for the observed functions of MAG (see below; Vinson et al., 2001). MAG has also been shown to interact with several collagens in the ECM (Fahrig et al., 1987), and recently, MAG was shown to interact with the Nogo‐66 receptor (> Table 2-33) (Domeniconi et al., 2002).

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. Table 2-33 Binding partners for MAG CAM MAG

Extracellular binding partner GD1a and GT1b Collagens Nogo‐66 receptor

Reference Yang et al. (1996) Fahrig et al. (1987) Domeniconi et al. (2002)

12.8 Signaling MAG seems to modulate intracellular signaling e.g., through trans‐interactions with Nogo‐66 (Barton et al., 2003). MAG and Nogo‐66 mediate inhibition of neurite outgrowth by activating the small GTPase RhoA and the RhoA‐associated kinase, ROCK. At the same time an inhibition of another small GTPAse, Rac1, is seen. The effects can be blocked by elevating the level of cAMP or by stimulating PKA (Bandtlow, 2003). MAG is believed to inhibit neurite outgrowth through effects on growth cones. Thus, an extracellular gradient of MAG can induce an elevation of intracellular Ca2þ, thereby modulating growth cone progression (Henley et al., 2004).

12.9 Function MAG is a bifunctional molecule whose function is dependent on the developmental stage. Thus, it promotes neurite outgrowth in juvenile neurons, but inhibits neurite extension from neurons in the mature nervous system (Skaper et al., 2001). The inhibitory properties of MAG are effectuated through growth cone collapse and are potent both in vitro and in vivo (Li et al., 1996). MAG is also important for the formation of myelin, and promotes myelination in the CNS. However, it does not seem to be essential for the formation of myelin in the PNS, where MAG knockout mice display a normal thickness of the myelin layer and timing of myelin formation (Li et al., 1994; Montag et al., 1994; Bartsch et al., 1997).

13

Neural Cell Adhesion Molecule

13.1 Introduction The neural cell adhesion molecule (NCAM) (> Table 2-34) was the first neural cell adhesion molecule to be identified (Jørgensen and Bock, 1974; Rutishauser et al., 1976), and the protein is also one of the most abundant adhesion proteins in the nervous system constituting around 3% of the total protein in the developing rat brain (Linnemann et al., 1993). . Table 2-34 Nomenclature for NCAM CAM name NCAM

Protein and gene synonyms CD56, D2, N‐CAM, NCAM1

13.2 Gene NCAM exists in three main isotypes obtained by alternative splicing of the transcript from a single gene (see > Table 2-35). Thus, mRNAs of 7.4 and 6.7 kb are translated into NCAM‐180 and NCAM‐140,


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. Table 2-35 Gene information for NCAM Species Man Mouse

Approved gene symbol NCAM1 Ncam1

Approved gene name Neural cell adhesion molecule 1

Locus 11q23–q24 9 28.0


respectively, whereas mRNAs of 5.2 and 2.9 kb are translated into NCAM‐120 (the difference in size of mRNA coding for NCAM‐120 is due to alternative polyadenylation (Linnemann et al., 1993). Transcription factors reported to be of importance for the regulation of NCAM expression includes Alx4, Barx2, Cux, Hoxb8, Hoxb9, Hoxc6, LEF‐1, NF‐kB, Otx2, Pax3, Pax6, Pax8, Phox2, and rhBMP‐2 (Jones et al., 1992; Valarche et al., 1993; Holst et al., 1994, 1997; Kioussi et al., 1995; Boersma et al., 1999; Nguyen Ba‐Charvet et al., 1999a; Edelman et al., 2000; Simpson and Morris, 2000; Boras and Hamel, 2002).

13.3 Expression NCAM is expressed in a developmentally regulated manner in many tissues including lungs (Filiz et al., 2002), skeletal and heart muscles (Burroughs et al., 1991; Reyes et al., 1991; Lyons et al., 1992; Andersson et al., 1993; Cifuentes‐Diaz et al., 1993; Gaardsvoll et al., 1993; Fidzianska and Kaminska, 1995), the male reproductive system (Møller et al., 1991; Li et al., 1998), the digestive system (Esni et al., 1999; Libbrecht et al., 2001), and in natural killer cells (Carson and Caligiuri, 1996). In the nervous system, NCAM is expressed by several cell types including neurons, astrocytes, oligodendrocytes, and Schwann cells, and its level of expression is isotype specific and temporally and spatially regulated (Theveniau et al., 1992; Katar et al., 1993; Linnemann et a., 1993; Martini, 1994). The individual isotypes of NCAM exhibit different subcellular localizations. Thus, NCAM‐180 is almost exclusively restricted to postsynaptic membranes in neurons, whereas NCAM‐140 is expressed on both pre‐ and postsynaptic membranes (Persohn et al., 1989; Schuster et al., 1998, 2001). N‐CAM‐120 is not detectable in synaptosomal membranes (Persohn et al., 1989) but is like many other GPI‐linked proteins predominantly found in detergent‐resistant microdomains (Kra¨mer et al., 1999).

13.4 Isoforms and Protein Structure The extracellular part of NCAM contains five Ig‐modules (denoted IgI‐V) followed by two Fn3‐modules (denoted Fn3I and Fn3II) (see > Figure 2-1). Ig‐I, ‐II, and III are of the intermediate type (Thomsen et al., 1996; Jensen et al., 1999; Soroka et al., 2003). NCAM‐180 and ‐140 are transmembrane isoforms, which only differ in the length of the cytoplasmic tail, whereas NCAM‐120 is attached to the membrane by a GPI anchor (the three subtypes are sometimes referred to as NCAM‐A, ‐B, and ‐C or ld, sd, and ssd, respectively). In the mouse, the gene consists of exons 0–19 plus 6 additional smaller exons. Exons 0–14 encode the extracellular part of the protein. Expression of exon 15 results in the production of the GPI‐linked isoform, whereas exon 16 encodes the transmembrane segment, and exon 17–19 encode the intracellular part of the molecule. Exon 18 is specific for the NCAM‐180 isoform (Cunningham et al., 1987). The many isoforms of NCAM are obtained by the exclusion or inclusion of six small exons in the original transcript. The smallest exon (positioned between exon 12 and exon 13) is named AAG and consists only of a single nucleotide triplet. The variable alternative‐spliced exon (VASE) or p‐exon (positioned between exon 7 and exon 8) encodes a 10‐amino acid‐long sequence known to have an inhibitory effect on neurite outgrowth (Doherty et al., 1992; Liu et al., 1993; Arce et al., 1996; Lahrtz et al., 1997), and whose expression may possibly be related to psychiatric disorders (Vawter, 2000). The three exons MSD1a, ‐b, and ‐c (positioned between exon 13 and exon 12) comprise the so‐called muscle specific domain 1, MSD1 (Dickson et al., 1987; Santoni et al., 1989; Thompson et al., 1989; Hamspere et al., 1991; Reyes et al., 1991; Barthels et al., 1992).

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The last small exon is the SEC exon (positioned between exon 13 and exon 12). This exon contains a stop codon, thereby producing a truncated form of the extracellular part of NCAM, which is secreted into the extracellular space, SEC‐NCAM (Gower et al., 1988). In addition, NCAM also exists in shed forms produced either by enzymatic removal of NCAM‐120 from the membrane by the phosphatidylinositol‐specific phospholipase C (PI‐PLC) or by proteolytic cleavage of the extracellular part of any of the three major subtypes (He et al., 1986; Nybroe et al., 1989; Diestel et al., 2005; Hubschmann et al., 2005). Furthermore, NCAM‐expressing cells shed soluble isoforms corresponding to intact NCAM‐140 and ‐180, and intact transmembrane isoforms of NCAM can be found in cerebrospinal fluid (Olsen et al., 1993).

13.5 Posttranslational Modifications The cytoplasmic tail of NCAM‐140 and ‐180 can be phosphorylated at serine and threonine residues (Annunziata et al., 1983; Gennarini et al., 1984; Lyles et al., 1984; Sorkin et al., 1984), and recently, human NCAM‐180 was reported to be tyrosine phosphorylated on Y734 (Diestel et al., 2004). The phosphorylation of at least one threonine residue has been shown to be of importance for NCAM‐mediated activation of the transcription factor NF‐kB (Little et al., 2001), whereas the tyrosine phosphorylation seems to have an inhibitory effect on neurite outgrowth (Diestel et al., 2004). The transmembrane isoforms of the protein are anchored to the plasma membrane by palmitoylation of 2–4 highly conserved cysteine residues (Little et al., 1998). Abrogation of palmitoylation affects the distribution of NCAM within the plasma membrane and disrupts NCAM‐mediated signaling and neurite outgrowth (Niethammer et al., 2002). NCAM contains six N‐glycosylation sites (Albach et al., 2004), whose glycosylation pattern is spatially and temporally regulated (Schwarting et al., 1987; Yoshihara et al., 1991; Sasaki and Endo, 1999). It is one of very few vertebrate proteins to be glycosylated with polysialic acid, PSA (large homopolymers of the negatively charged molecule a2,8‐sialic acid) (Hoffman et al., 1982; Finne et al., 1983; McCoy et al., 1985), which is attached to two N‐glycosylation sites in the IgV‐module (Angata et al., 1995; Nelson et al., 1995; Liedtke et al., 2001; von Der Ohe et al., 2002; Wuhrer et al., 2003). NCAM also expresses the carbohydrate epitope HNK‐1, which can be attached to NCAM by N‐linked glycosylation at five locations, the actual glycosylation pattern varying between species (Crossin et al., 1984; Lyles et al., 1984; Liedtke et al., 2001; Wuhrer et al., 2003; Albach et al., 2004). In addition, NCAM can express O‐linked HNK‐1 epitopes attached to the MSD1‐region (Walsh et al., 1989; Ong et al., 2002), and the protein can also contain a number of non‐PSA/non‐HNK‐1 glycosylations, of which several are specific for the olfactory system (Key and Akeson, 1990, 1991; Pestean et al., 1995; Dowsing et al., 1997). For reviews of glycosylation of neural Ig CAMs see Durbec and Cremer (2001), Krog and Bock (1992), Kiss and Rougon (1997), and Kleene and Schachner (2004).

13.6 Homophilic Interactions NCAM is involved in homophilic cis‐ and trans‐interactions (> Table 2-36). It has been proposed that the homophilic trans‐interaction involved all five Ig‐modules, and that IgI, IgII, IgIII, IgIV and IgV of one NCAM molecule bound to IgV, IgIV, IgIII, IgII and IgI, respectively, of another NCAM molecule (Rao et al., 1992; Ranheim et al., 1996). However, data derived from structures based on nuclear magnetic resonance (NMR)‐ or X‐ray crystallography of up to three‐modules‐long recombinant constructs do not support this model (Thomsen et al., 1996; Atkins et al., 1999, 2001; Jensen et al., 1999; Kasper et al., 2000; Soroka et al., 2003; Kiselyov et al., 2005). A recent crystal structure of the IgI–IgII–IgIII triple module of NCAM reveals four intermodular interactions. Three of these interactions occur between antiparallel triple modules and are therefore believed to represent homophilic trans‐interaction, whereas the fourth interaction is believed to represent a homophilic cis‐interaction. The proposed trans‐interactions include IgII binding to IgIII, IgI binding to IgIII, and IgII binding to IgII of an NCAM molecule on an opposing cell surface. The proposed cis‐interaction is formed by the binding of the IgI and IgII modules of one NCAM molecule to the IgII and


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. Table 2-36 Binding partners for NCAM Extracellular binding partners NCAM (cis, trans) ATP Collagen CSPGs: neurocan, phosphacan FGFR GDNF and GFRa Heparin HSPGs: agrin and 6C4 L1 Laminin P‐ and L‐selectin TAG‐1 Spectrin b‐Actin, a‐actinin, a/b‐tubulin, MAP1A, tropomyosin, ROKa LANP, PACSIN1, PLCg, PP1, PP2A, TOAD‐64 PKCb Fyn FAK GAP43 RPTPa

Reference Soroka et al. (2003) Dzhandzhugazyan and Bock (1993, 1997); Kiselyov et al. (2003) Kiselyov et al. (1997); Probstmeier et al. (1989, 1992) Grumet et al. (1993); Friedlander et al. (1994); Retzler et al. (1996); Milev et al. (1994, 1995) Kiselyov et al. (2003); Williams et al. (1994a) Paratcha et al. (2003) Cole and Akeson (1989) Burg et al. (1995); Herndon et al. (1999); Storms et al. (1996b) Storm and Rutishauser (1998); Storms et al. (1996a) Heiland et al. (1998); Horstkorte et al. (1993) Hall et al. (1993) Needham and Schnaar (1993) Milev et al. (1996) Pollerberg et al. (1986, 1987); Leshchyns’ka et al. (2003) Bu¨ttner et al. (2003) Bu¨ttner et al. (2005) Leshchyns’ka et al. (2003) Beggs et al. (1997); Kra¨mer et al. (1999) Beggs et al. (1997) He and Meiri (2002) Bodrikov et al. (2005)

IgI modules, respectively, of another NCAM molecule on the same cell surface (Kiselyov et al., 1997, 2005; Atkins et al., 1999; Jensen et al., 1999; Kasper et al., 2000; Soroka, et al., 2003). Recently, a ‘‘double zipper model,’’ which involves the combination of homophilic trans‐ and cis‐ interactions, was proposed. A hypothetical ‘‘flat zipper’’ is formed between NCAM cis‐dimers on one cell surface interacting in trans through IgII–IgIII contacts with NCAM cis‐dimers on another cell surface. A ‘‘compact zipper’’ is formed between NCAM cis‐dimers on one cell surface interacting in trans through IgI–IgIII and IgII–IgII contacts with NCAM cis‐dimers on another cell surface. The two zippers can theoretically be formed perpendicularly to each other. It has therefore been proposed that homophilic NCAM cis‐dimerization subsequently may lead to the formation of ‘‘compact flat double zippers,’’ thereby potentially producing homophilic NCAM adhesion complexes involving numerous NCAM molecules (> Figure 2-7) (Soroka et al., 2003; Kiselyov et al., 2005).

13.7 Heterophilic Binding Partners 13.7.1 Extracellular Binding Partners NCAM binds to the related adhesion molecule L1 (> Table 2-36). The interaction probably occurs as a cis‐ interaction between carbohydrates expressed on L1 and a lectin homology motif in the IgIV module of NCAM (Horstkorte et al., 1993; Heiland et al., 1998). The interaction seems to be dependent on simultaneous NCAM–NCAM interactions and has synergistic effects on L1‐mediated cellular aggregation and

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. Figure 2-7 Homophilic NCAM Zippers (a): Crystral structure of NCAM Ig1–Ig2–Ig3. The figure shows interactions between the three Ig‐modules resulting in a hypothetical flat zipper (left) or compact zipper (right). (b): Illustration of the organization of the extracellular part of NCAM molecules engaged in a flat zipper (left) and compact zipper (right). (c): Illustration of the organization of the extracellular part of NCAM molecules engaged in a both a flat and a compact zipper. The large ellipsoids in (b) and (c) correspond to two interacting Ig1–Ig2–Ig3 constructs as shown in the leftmost part of (a) (Modified from Kiselyov et al., 2005)

adhesion, a phenomenon, which has been termed ‘‘assisted homophilic L1–L1 trans‐binding’’ (Kadmon et al., 1990a, b; Kristiansen et al., 1999). NCAM also binds to the GPI‐anchored adhesion molecule TAG‐1 (Milev et al., 1996). TAG‐1 also binds to L1 in a cis‐interaction (Malhotra et al., 1998), thereby making an L1/NCAM/TAG‐1 complex theoretically possible. Alternatively, L1/TAG‐1 and NCAM/TAG‐1 may be mutually exclusive. NCAM has ecto‐ATPase activity. The ATP‐binding site is located in the Fn3II module of NCAM (Kiselyov et al., 2003), and studies suggest that binding of ATP to NCAM inhibits cellular aggregation and neurite outgrowth by interfering with homophilic NCAM interactions and signaling (Dzhandzhugazyan and Bock, 1993, 1997; Skladchikova et al., 1999). The Fn3II module of NCAM interacts in cis with the D3 module of the fibroblast growth factor receptor, FGFR. The ATP‐binding site on NCAM overlaps with the FGFR binding site, and ATP can inhibit NCAM‐induced signaling mediated through FGFR (Williams et al., 1994a; Kiselyov et al., 2003). NCAM binds directly to GDNF and the GPI‐linked GFRa receptor. The binding of GDNF does not affect homophilic NCAM interactions, whereas binding of GFRa to NCAM abrogates homophilic NCAM


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trans‐interactions, but at the same time potentates the GDNF–NCAM interaction. The reported effects of the NCAM/GFRa/GDNF interactions include the induction of neurite outgrowth and Schwann cell migration (Paratcha et al., 2003), but recent in vivo studies indicate that GFRa‐induced NCAM signaling only plays a minor physiological role (Enomoto et al., 2004). NCAM interacts with the ECM components heparin, collagen, and laminin as well as several chondroitin sulfate proteoglycans (CSPGs) and heparan sulfate proteoglycans (HSPGs), (e.g., agrin, neurocan, and phosphacan) (Grumet et al., 1993; Burg et al., 1995; Milev et al., 1995; Storms et al., 1996a, b; Herndon et al., 1999). Heparin binds to a so‐called ‘‘heparin binding domain,’’ in the IgII module of NCAM (Cole and Akeson, 1989; Kulahin et al., 2005), but the interaction may also involve the IgI module of NCAM (Kiselyov et al., 1997). The binding to collagen (I–IV and IX) may be indirect via heparan sulfate (Probstmeier et al., 1989, 1992; Kiselyov et al., 1997), whereas interactions with laminin as well as putative interactions with P‐ and L‐selectin supposedly are mediated by HNK‐1‐epitope carrying carbohydrates on NCAM (Hall et al., 1993; Needham and Schnaar, 1993). Agrin has been suggested to interact with NCAM both at the heparin‐binding domain and via PSA on the IgV‐module (Storm and Rutishauser, 1998). Phosphacan is an alternative‐splicing product representing the extracellular part of the receptor‐like protein tyrosine phosphatase (RPTP) x/b. NCAM is believed to interact with N‐linked carbohydrates on phosphacan via the lectin homology motif in the IgIV module (Horstkorte et al., 1993; Milev et al., 1995). NCAM– neurocan interactions interfere with homophilic NCAM interactions and inhibit neuronal adhesion and neurite outgrowth (Friedlander et al., 1994; Retzler et al., 1996). Finally, NCAM is a receptor for rabies virus and for the cellular prion protein. The virus may bind to the IgI and/or IgII module, whereas the prion protein is thought to bind NCAM at the IgV, Fn3I and/or Fn3II modules (Thoulouze et al., 1998; Schmitt‐Ulms et al., 2001) (see also > Table 2-36).

13.7.2 Intracellular Binding Partners Proteins that have been reported to interact with NCAM include ROKa and the cytoskeletal proteins spectrin, a‐ and b‐tubulin, a‐actinin, MAP1A, b‐actin, and tropomyosin, as observed in immunoprecipitation experiments (Pollerberg et al., 1986, 1987; Bu¨ttner et al., 2003; Leshchyns’ka et al., 2003) (> Table 2-36). Furthermore, using pull‐down assays, the following proteins were recently found to bind the cytoplasmic tail of NCAM‐140 and/or ‐180: PLCg, LANP (a phosphatase inhibitor), PACSIN1/syndapin (a protein involved in vesicle trafficking), TOAD‐64 (a protein involved in axonal growth), and the serine/threonine phosphatases PP1 and PP2A. PLCg, PACSIN1, PP1, PP2A, TOAD‐64, and a and b tubulin have been found to bind to the membrane proximal region of the cytoplasmic tail (encoded by exon 16–17). ROKa binds to the NCAM‐180‐specific region encoded by exon 18, whereas LANP binds to the membrane distal C‐terminal encoded by exon 19 (Buttner et al., 2005). Additional signaling molecules reported to interact with NCAM include PKCb (Leshchyns’ka et al., 2003), FAK, GAP43, and the Src family nonreceptor tyrosine kinase Fyn. The interaction with FAK is believed to occur indirectly via Fyn, which may be activated through a direct interaction between NCAM and RPTPa (Beggs et al., 1997; He and Meiri, 2002; Bodrikov et al., 2005) (see also > Table 2-36). Some of the intracellular interactions of NCAM may occur at the sequence TEVKT (amino acid 839– 843 in exon 19 of rat NCAM), which has been shown to be of importance for neuritogenesis (Kolkova et al., 2000a).

13.8 Signaling Homophilic and heterophilic NCAM interactions initiate a number of intracellular signaling pathways. The exact signaling molecules involved in NCAM‐mediated signaling are to some extent cell‐type specific, and also depend on the type of event promoted by the signal (cellular survival, differentiation etc.). However, the description given below only serves to present the general pathways involved in NCAM signaling and does not discriminate between cell types or signal outcome.

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Cross‐linking of NCAM molecules or interactions between NCAM, GDNF, and GFRa can induce phosphorylation of the nonreceptor tyrosine kinases Fyn and FAK, subsequently leading to stimulation of the MAPK pathway (Beggs et al., 1997; Saffell et al., 1997; Paratcha et al., 2003). FAK interacts with numerous proteins and may activate the MAPK pathway via several routes (Barberis et al., 2000; Schaller, 2001), but ultimately the induction of this pathway leads to activation of the kinases MEK1/2 and ERK1/2 and of the transcription factors CREB and fos (Deak et al., 1998; Jessen et al., 2001; Schmid et al., 1999). Extracellular interactions between NCAM and FGFR induce dimerization and autophosphorylation of FGFR (Kiselyov et al., 2003). Subsequently, the cytoplasmic tail of FGFR serves as docking site for several proteins including PLCg, which becomes activated upon binding. PLCg cleaves phosphatidylinositol‐4,5‐ bisphosphate (PIP2) to generate inositol‐1,4,5‐trisphosphate (IP3) and diacylglycerol (DAG). IP3 binds to intracellular Ca2þ channels, thereby inducing the release of Ca2þ from intracellular stores, whereas DAG either can activate PKC (see below) or can be converted to 2‐arachidonylglycerol (2‐AG) and arachidonic acid (AA), which both can activate calcium influx via calcium channels in the plasma membrane (Williams et al., 1994b, 2003; Sugiura and Waku, 2002; Shuttleworth and Mignen, 2003). One effect of the NCAM‐ mediated increase in intracellular calcium concentrations is activation of CaMKII (Williams et al., 1995), which subsequently may activate CREB. Interestingly, the intracellular signal transduction induced by NCAM‐stimulated activation of FGFR is different when compared to activation of FGFR by FGF2. For instance, the docking protein ShcA is important for NCAM‐induced, but not FGF2‐induced neurite outgrowth mediated through FGFR (Hinsby et al., 2004b). NCAM can activate PKC either via FGFR (see above) or via spectrin (Leshchyns’ka et al., 2003) leading to the activation of GAP‐43 (Meiri et al., 1998) and c‐Raf‐1 (and thereby the MAPK pathway) (Kolkova et al., 2000b). Recent investigations have demonstrated that at least PKCa, bI, bII, and e are important for NCAM‐mediated neurite outgrowth (Kolkova et al., 2005). NCAM can also activate heterotrimeric G proteins (Doherty et al., 1991; Lipkin et al., 1992; Sandig et al., 1994) leading to stimulation of adenylyl cyclase and production cAMP, which subsequently leads to the activation of PKA and ultimately induces the activation of CREB and another transcription factor, c‐Fos (Jessen et al., 2001). NCAM can also activate PI3K and PKB (Ditlevsen et al., 2003). PI3K may be activated via either FGFR, FAK, or Ras (Nguyen et al., 1993; Rodriguez‐Viciana et al., 1994; Ong et al., 2001; Steelman et al., 2004), and subsequently, PKB may be activated through phosphorylation by PDK after binding the lipid products of PI3K (Ong et al., 2001; Steelman et al., 2004). The targets downstream of PKB may include transcription factors such as CREB and NF‐kB (Du and Montminy, 1998; Kane et al. 1999; Krushel et al., 1999; Choi et al., 2001).

13.9 Functions NCAM is involved in numerous physiological phenomena including embryonic development, neuronal regeneration, and plasticity (learning and memory formation) through its effects on cell–cell and cell–ECM interactions, cellular proliferation (Mistry et al., 2002), differentiation (Nguyen et al., 2003), survival (Ditlevsen et al., 2003; Pedersen et al., 2004), motility (Prag et al., 2002), axon guidance (Walsh and Doherty, 1997), and synaptic plasticity in vitro and in vivo (Welzl and Stork, 2003; Yamagata et al., 2003; Cambon et al., 2004). Some of the effects of NCAM are dependent on or modulated by the expression of PSA. For instance, BDNF‐mediated differentiation and survival of cortical neurons is reduced when the cells are stripped for PSA or cultured in the presence of excess free PSA (Vutskits et al., 2001), and the migration, proliferation, and adhesion of cells are also affected by the removal of PSA (Seidenfaden et al., 2003; Zhang et al., 2004). Furthermore, PSA is expressed predominantly on NCAM during embryonic development, and PSA– NCAM is therefore sometimes referred to as ‘‘embryonic’’‐ or E‐NCAM, as opposed to NCAM containing little or no PSA referred to as ‘‘adult’’‐ or A‐NCAM (Edelman and Choung, 1982). However, in areas whose functions require neural plasticity (such as the hippocampus, the olfactory system, and the hypophysis), PSA is expressed throughout life (Kiss and Rougon, 1997; Cremer et al., 2000; Bruses and Rutishauser, 2001; Durbec and Cremer, 2001).


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Deletion of NCAM is not lethal. Thus, NCAM‐deficient mice are fertile and appear healthy. However, they exhibit 10% decrease in brain weight, and demonstrate alterations in the olfactory bulb and hippocampus. The alterations in the hippocampus are more severe in adult mice than in young mice, indicating that NCAM is important throughout life, for the maintenance of the nervous system (Cremer et al., 1994, 1997). Behaviorally, the activity and motoric abilities of NCAM‐deficient mice appear normal. However, the animals demonstrate defects in spatial learning, and exhibit an increased intermale aggressivity and anxiety‐ like behaviors. Re‐expression of NCAM‐180 in NCAM‐deficient mice can counteract some of the behavioral alterations observed in response to the lack of NCAM, but cannot improve the observed reduction in learning (Stork et al., 1997, 1999, 2000). Expression of soluble, secreted NCAM in mice not expressing membrane‐associated NCAM has been demonstrated to be embryolethal, demonstrating that the extracellular region of NCAM induces signaling via extracellular heterophilic ligands (Rabinowitz et al., 1996). Furthermore, the simultaneous deletion of the two genes encoding the polysialyltransferases responsible for the polysialylation of NCAM, St8sia2, and St8sia4 results in a severe phenotype, which includes postnatal growth retardation and early death (Weinhold et al., 2005). For recent reviews of NCAM see Povlsen et al. (2003); Hinsby et al. (2004a); Walmod et al. (2004); Kiselyov et al. (2005).

14

Neural Cell Adhesion Molecule 2

14.1 Introduction NCAM2 was originally described in 1986 as a membrane associated 125‐kDa protein identified by the monoclonal antibody Rb‐8 (Schwob and Gottlieb, 1986, 1988). In 1997, three groups independently cloned the protein from humans, mice, rats, and rabbits. The protein identified in humans was named NCAM2 (Paoloni‐Giacobino et al., 1997), whereas the proteins identified in mice, rats, and rabbits were named RNCAM (Rb‐8‐neural cell adhesion molecule) (Alenius and Bohm, 1997) and OCAM (olfactory cell adhesion molecule) (Yoshihara et al., 1997) (> Table 2-37). . Table 2-37 Nomenclature for NCAM2 CAM name NCAM2

Protein and gene synonyms mamFas II, N‐CAM2, neural cell adhesion molecule 2, OCAM, R4B12 antigen, RB‐8, RNCAM

14.2 Gene NCAM2 mRNA exists in several sizes obtained by alternative splicing of the transcript from a single gene. The untranslated 30 region of NCAM2 mRNAs contains a number of so‐called ATTTA motifs, believed to confer instability to the mRNA. This motif is present in largest numbers in mRNA encoding GPI‐linked NCAM2 (Yoshihara et al., 1997) (see also > Table 2-38). . Table 2-38 Gene information for NCAM2 Species Man Mouse

Approved gene symbol NCAM2 Ncam2

Approved gene name Neural cell adhesion molecule 2

Locus 21q21 16 56.0 C1–3


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14.3 Expression In humans, Northern blots have identified NCAM2 transcripts in all identified brain regions, and in several other tissues including lung, kidney, heart, and testis (Paoloni‐Giacombino et al., 1997). In mouse, NCAM2 is almost exclusively expressed in neuronal tissue and predominantly in the olfactory sensory system (Alenius and Bohm, 1997; Yoshihara et al., 1997). More specifically, NCAM2 is expressed on sensory neurons in zones II, III, and IV (but not zone I) of the olfactory epithelium (according to the terminology of Ressler et al., 1993), in sensory neurons of the vomeronasal organ expressing Gai2 G proteins (Alenius and Bohm, 1997; Yoshihara et al., 1997), and on the dendrites of mitral/tufted (M/T) cells in the accessory olfactory bulb (von Campenhausen et al., 1997; Treloar et al., 2003). In the mouse brain, the expression level of the transmembrane isoform of NCAM2 is generally higher than that of GPI‐anchored NCAM2. The protein is detectable at E18, peaks at P(21), but maintains a steady expression level in the adult mouse (Yoshihara et al., 1997).

14.4 Isoforms and Protein Structure The overall structure of NCAM2 is similar to that of NCAM. Thus, NCAM2 consists of five Ig‐modules (supposedly of the C2‐type) followed by two Fn3‐modules. The protein exists in a GPI‐anchored isoform and in a transmembrane isoform that contains a 20–25 amino acid transmembrane segment followed by a 106–119‐amino acid cytoplasmic tail (Alenius and Bohm, 1997; Yoshihara et al., 1997). Thus, the two isoforms of NCAM2 are comparable to NCAM‐120 and NCAM‐140. The overall amino acid identity between NCAM and NCAM2 is around 46% for the respective Ig‐modules, around 37% for the Fn‐III‐modules, and around 54% for the cytoplasmic tail (Alenius and Bohm, 1997; Paoloni‐ Giacobino et al., 1997; Yoshihara et al., 1997). The transmembrane isoform of NCAM2 contains a PEST sequence in the cytoplasmic tail that makes the protein susceptible to proteolytic degradation (Yoshihara et al., 1997).

14.5 Posttranslational Modifications NCAM2 contains 8–9 potential sites for N‐linked glycosylation, of which some correspond to the PSA‐ glycosylated residues on NCAM (see Sect. 15). However, whereas NCAM2 has been reported to bear HNK‐1 carbohydrate epitopes, it has not been demonstrated to be glycosylated by PSA (Paoloni‐Giacobino et al., 1997; Yoshihara et al., 1997).

14.6 Homophilic Interactions Experiments have demonstrated that fibroblasts transfected with NCAM2 aggregate more strongly than the parental nontransfected cells, indicating that NCAM2 interacts homophilically in trans. Data from in vitro binding assays support this notion, but the modules involved in the homophilic interactions are not identified, neither has it been demonstrated whether NCAM2—like NCAM—is involved in homophilic cis‐interactions (Yoshihara et al., 1997) (> Table 2-39). . Table 2-39 Binding partners for NCAM2 Extracellular binding partners NCAM2 (trans)

Reference Yoshihara et al. (1997)


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14.7 Heterophilic Binding Partners NCAM2 has not yet been demonstrated to be involved in any heterophilic extracellular or intracellular interactions. However, it has been demonstrated that the protein does not bind to NCAM (Yoshihara et al., 1997).

14.8 Functions NCAM2 appears to be involved in neurite outgrowth, axonal guidance, and synapse formation in the olfactory system. Thus, experiments have demonstrated a correlation between the expression of NCAM2 on olfactory neurons and their axonal growth and fasciculation in vitro (Hamlin et al., 2004). Furthermore, axons of NCAM2‐negative neurons in the olfactory and vomeronasal epithelium have been reported to form synapses with NCAM2‐positive M/T‐cells, whereas axons of NCAM2‐positive neurons in the olfactory and vomeronasal epithelium form synapses with NCAM2‐negative M/T‐cells (von Campenhausen et al., 1997; Treloar et al., 2003). Furthermore, overexpression of GPI and/or transmembrane NCAM2 has been reported to interfere with the axonal guidance to defined axon convergence sites (the glomeruli) in the olfactory bulb in an isotype‐specific manner (Alenius and Bohm, 2003). In rats, NCAM2 has been described to be of importance for the formation of dendritic bundles in brain regions involved in spatial learning and memory (Ichinohe et al., 2003). In humans, the NCAM2 gene is located on chromosome 21. Down syndrome is characterized by trisomy 21, and consequently it has been proposed that a potentially excessive production of NCAM2 might be a contributing factor to the symptoms of the disease (Paoloni‐Giacobino et al., 1997). However, a mouse model of Down syndrome, which confers trisomy of a distal part of chromosome 16, was shown not to include the NCAM2 gene, although the locus for NCAM2 in mouse is present on chromosome 16 (Akeson et al., 2001).

15

The Robo Family

15.1 Introduction The Robo family constitutes a part of a larger group of IGSF proteins referred to as molecular guidance molecules (> Table 2-40). Originally, the protein Robo was identified in Drosophila, and several of the major studies of Robo function have been performed in this organism.

. Table 2-40 Nomenclature for Robo family members CAM name ROBO1 ROBO2 ROBO3 ROBO4

Protein and gene synonyms DUTT1, FLJ21882, roundabout, axon guidance receptor homolog 1, SAX3 2600013A04Rik, 9430089E08Rik, D230004I22Rik, KIAA1568, roundabout, axon guidance receptor homolog 2 FLJ21044, Rbig1, RBIG1, Rig‐1, roundabout axon guidance receptor homolog 3 1200012D01Rik, FLJ20798, magic roundabout, roundabout homolog 4

The mammalian Robo family constitutes four members: Robo1 and Robo2, which were cloned in 1998 (Kidd et al., 1998; Sundaresan et al., 1998a, b), Robo3, which was cloned the year after (Yuan et al., 1999a), and Robo4, which was cloned in 2002 (Huminiecki et al., 2002) (> Table 2-41).

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. Table 2-41 Gene information for Robo family members Species Man

Approved gene symbol ROBO1 ROBO2 ROBO3 ROBO4

Mouse

Robo1 Robo2 Robo3 Robo4

Approved gene name roundabout, axon guidance receptor, homolog 1 (Drosophila) roundabout, axon guidance receptor, homolog 2 (Drosophila) roundabout, axon guidance receptor, homolog 3 (Drosophila) roundabout homolog 4, magic roundabout (Drosophila) roundabout homolog 1 (Drosophila) roundabout homolog 2 (Drosophila) roundabout homolog 3 (Drosophila) roundabout homolog 4 (Drosophila)

Locus 3p12.3


3p12.3

HGNC ID: 10250

11q24

HGNC ID: 13433

11

HGNC ID: 17985

16 syntenic 16 syntenic 9B 9 syntenic

MGI ID: 1274781 MGI ID: 1890110 MGI ID: 1343102 MGI ID: 1921394

15.2 Genes There are indications for alternative splicing of the transcripts for Robo1 and Robo2 (Kidd et al., 1998), and in the transcripts for Robo3, at least nine alternatively spliced exons have been identified, the longest transcript consisting of 28 exons (Yuan et al., 1999a; Jen et al., 2004). Human ROBO3 consists of 18 exons (Huminiecki et al., 2002), and this protein also seems to exist in multiple splice variants. The promoter region of Robo3 is regulated in a cell type‐specific manner. The expression of the receptor is upregulated by the transcription factor Pax2, but downregulated by Rb, which affects transcription through a direct interaction with Pax2. At E11.5 of mouse development, Pax2 and Robo3 are coexpressed in certain regions of the CNS (Yuan et al., 2002).

15.3 Expression In general, Robo1 is ubiquitously expressed (Sundaresan et al., 1998a). In the nervous system, Robo1, Robo2, and Robo3 are found in the hindbrain and spinal cord, where their expressions are partially overlapping (Camurri et al., 2004). The expression patterns of Robo proteins in the CNS have been examined in detail in several studies (Sundaresan et al., 1998a, 2004; Nguyen Ba‐Charvet et al., 1999b, 2004; Yuan et al., 1999b; Erskine et al., 2000; Ringstedt et al., 2000; Marillat et al., 2002; Camurri et al., 2004). In the mouse, Robo3 is exclusively expressed in the nervous system where it is transiently expressed, the amount peaking around E11.5–12.5 (Yuan et al., 1999a). Robo4 is exclusively expressed in endothelial cells at sites of active angiogenesis (Huminiecki et al., 2002).

15.4 Isoforms and Primary Protein Structure Robo1, ‐2, and ‐3 consist of five Ig‐modules followed by three membrane‐proximal Fn3‐modules, a transmembrane segment, and a cytoplasmic tail (up to 700 amino acids) (Sundaresan et al., 1998a; Yuan et al., 1999a). Robo4 contains only two Ig‐modules and two Fn3‐modules (corresponding to the two most membrane‐distal Ig‐modules and the two most membrane‐proximal Fn3‐modules of Robo1) (Huminiecki et al., 2002).


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Within the Robo family, the first and second Ig‐modules are the most conserved. In contrast, the cytoplasmic tail is highly variable, except for four conserved motifs, denoted CC0, CC1, CC2, and CC3. However, Robo3 does not contain CC1 (Kidd et al., 1998; Jen et al., 2004). Because of the alternative splicing of the transcripts at least Robo1 and Robo3 are believed to exist in secreted as well as transmembrane forms (Yuan et al., 1999a; Clark et al., 2002).

15.5 Posttranslational Modifications Robo proteins contain 4–12 potential sites for N‐linked glycosylation.

15.6 Homophilic Interactions Robo1 and Robo2 interact homophilically in trans, and these two proteins are also involved in heterophilic trans‐interactions with each other (Hivert et al., 2002) (> Table 2-42). However, it is unclear which of the EC modules mediate the interactions (Liu et al., 2004b).

. Table 2-42 Binding partners for Robo family members CAM Robo1

Robo2

Robo3 Robo4 Robo1

Robo4

Extracellular binding partner Robo1 (trans) Robo2 (trans) Slit1, Slit2, Slit3 Robo1 (trans) Robo2 (trans) Slit1, Slit2, Slit3 Slit2 Slit2 srGAP1, srGAP2, srGAP3 Vilse Dock Abl DCC Ena/VASP‐like protein

Reference Hivert et al. (2002) Hivert et al. (2002) Brose et al. (1999); Liu et al. (2004b); Suchting et al. (2004) Hivert et al. (2002) Hivert et al. (2002) Brose et al. (1999); Liu et al. (2004b) Sabatier et al. (2004) Park et al. (2003) Wong et al. (2001) Lundstrom et al. (2004) Fan et al. (2003) Bashaw et al. (2000) Stein and Tessier‐Lavigne (2001) Park et al. (2003)

15.7 Heterophilic Binding Partners 15.7.1 Extracellular Binding Partners Robo proteins interact extracellularly with the soluble protein Slit (> Table 2-42). In mammals, three Slit proteins have been identified (Slit1, Slit2, and Slit3). These 1521–1531‐amino acid‐long proteins consist of four leucine‐rich repeats, nine EGF‐like sequences, a laminin‐G domain containing agrin, laminin, and perlecan homologies (termed ALPs), and a C‐terminal cysteine‐rich region (Yuan et al., 1999b; Park et al., 2003). Slit2 binds to Robo1 and Robo2 with a Kd around 4–5 nM (Brose et al., 1999). The interaction with Slit is mediated by the Ig1‐ and Ig2‐module of Robo (Liu et al., 2004b), and the affinity between Slit2 and Robo1 has been shown to be increased in the presence of heparan sulfate (Hu, 2001). (See also review by Nguyen‐Ba‐Charvat and Che´dotal (2002)).

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Recently, it was proposed from studies made in Drosophila that Slit is included in a ‘‘Slit sandwich,’’ where the protein serves as a ‘‘linker’’ mediating Robo2 trans‐interactions (Kraut and Zinn, 2004). A recent study could not demonstrate interactions between Robo4 and any of the Slit proteins, whereas an earlier study has demonstrated an interaction between Robo4 and Slit2 (Park et al., 2003; Suchting et al., 2004).

15.7.2 Intracellular Binding Partners The intracellular tails of Robo proteins interact with a novel family of Rho GAPs referred to as Slit‐Robo GAPs (srGAPS). This family includes three proteins: srGAP1, srGAP2, and srGAP3/formin binding protein 2 (Wong et al., 2001). In addition, the CC2 region of Robo1 has been shown to interact with another GAP‐ protein, Vilse (KIAA1688 protein) (Lundstrom et al., 2004) (> Table 2-42). Robo1 has also been shown to bind to the adaptor protein Dock, the Src homology 3 (SH3) domain of the tyrosine kinase Abl, and the P3 domain of the receptor DCC (Bashaw et al., 2000; Stein and Tessier‐ Lavigne, 2001; Rhee et al., 2002; Fan et al., 2003). Robo4 binds to the protein Ena/VASP‐like protein, which is believed to affect the dynamics of the actin cytoskeleton (Park et al., 2003) (> Table 2-42). In Drosophila, the GTPase‐activating protein (GAP) CrossGAP/Vilse, which regulates the activity of the GTPase Rac, has been demonstrated to bind to Robo (Hu et al., 2005).

15.8 Signaling The interaction between Slit and Robo1 leads to an increase in the interaction between Robo1 and srGAP1. This subsequently leads to an inactivation of Cdc42, and coexpression experiments have shown that srGAP1 also can inactivate RhoA, but not Rac1 (Wong et al., 2001). The interaction between Robo1 and the GAP‐protein Vilse also leads to an inactivation of Rac and—to a smaller extent—Cdc42 (Lundstrom et al., 2004). In contrast, Robo1 in Drosophila has been shown to interact with the adaptor protein Dock, which can recruit the serine/threonine kinase Pak, and thereby stimulate Rac1 activity. The Robo1–Dock interaction is also enhanced by Robo–Slit interactions (Fan et al., 2003). See Ghose and van Vactor (2002) for a review of GAPs in Robo signaling. Slit–Robo interactions have been shown to inhibit cell adhesion mediated by N‐cadherin. This effect is possibly mediated through the Abl protein tyrosine kinase, which can interact with both Robo and N‐cadherin (Rhee et al., 2002). Recently, it was found that the effects of Slit on Robo1‐expressing cells can be reduced by expression of Robo3 (Sabatier et al., 2004).

15.9 Functions Robo proteins are molecular guidance molecules involved in axonal guidance and growth as well as regulation of cell migration. Slit is referred to as a repulsive guidance factor, and the expression of Slit proteins, for e.g., at the ventral midline, ensures that Robo‐expressing neurons do not cross or re‐cross the midline. Thus, human Slit2 collapses and repels retinal ganglion cell axons in vitro (Niclou et al., 2000), and if Slit expression is abrogated, many commissural axons either fail to leave the midline or re‐cross the midline during development. Furthermore, the knockout of either Robo1 or Robo2 also affects axonal guidance (Long et al., 2004), and Robo3 also controls midline crossing of axons (Marillat et al., 2004). (See Guthrie (2001) for a review). In the subventricular zone (SVZ), where both Robo2 and Robo3 and Slit 1 and Slit2 are expressed, the presence of Slit has been shown to be important for the migration of SVZ cells along the rostral migratory stream (Nguyen‐Ba‐Charvet et al., 2004).


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In addition to the regulation of axon guidance, Robos also regulate dendritic development. Thus, exposure to Slit1 can induce increased growth and branching of dendrites, and inhibition of Robo–Slit interactions leads to a reduction in the branching of dendrites (Whitford et al., 2002; Furrer et al., 2003). Furthermore, Robo1 and Robo2 can stimulate neurite outgrowth in Robo‐positive, but not in Robo‐ negative, neurons (Hivert et al., 2002). Mutations in Robo3 are related to a rare disease, horizontal gaze palsy with progressive scoliosis (HGPPS), which is characterized by the absence of coordinated horizontal eye movements (Jen et al., 2004) (see review by Guthrie, 2004).

16

The Contactin Family

16.1 Introduction The contactin family (F3, TAG‐1, contactins‐3 to ‐6) constitutes a group of proteins that all are composed of six Ig‐modules and four Fn3‐modules attached to the membrane by a GPI anchor. F3 was cloned in 1989 (Gennarini et al., 1989), the same year as the chicken orthologue F11 (Brummendorf et al., 1989). TAG‐1 was cloned the following year (Furley et al., 1990), and the related proteins, contactins‐3 to ‐6, were cloned a few years later (> Table 2-43) (Connelly et al., 1994; Yoshihara et al., 1994, 1995; Ogawa et al., 1996). All proteins of the contactin family are predominantly expressed in neural tissues. . Table 2-43 Nomenclature for contactin family members CAM name Contactin 3 Contactin 4 Contactin 5 Contactin 6 F3 TAG‐1

Protein and gene synonyms BIG‐1, PANG (plasmacytoma‐associated neuronal glycoprotein) Axcam, BIG‐2 hNB‐2, NB‐2 NB‐3 CNTN, contactin 1, F3cam Axonin, AXT, contactin 2, D130012K04Rik, TAX1

16.2 F3 16.2.1 Genes The promoter region of mouse F3 has been found to include both positive and negative regulatory elements, including a neural‐specific enhancer element. Factors regulating F3 expression include cAMP and retinoic acid (Cangiano et al., 1997). The mouse and rat genes encoding F3 are differently organized, and the rat gene contains multiple transcription initiation sites and two untranslated exons (Rome et al., 2005) (> Table 2-44).

16.2.2 Expression F3 is mainly expressed in the parallel fibers and postmitotic neurons of the cerebellum, in the dorsal root ganglion, and in the hippocampus (Yoshihara et al., 1995). A detailed study of the expression of F3 during mouse development demonstrated a regulation of the expression at the regional, cellular, and subcellular levels. For instance, the proteins shifts from being expressed on both cell bodies and processes early in development to predominantly being expressed on neurites later in development (Virgintino et al., 1999). In the adult rat, a decline in the expression of F3 can be observed in the hippocampus of 30‐month‐old

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. Table 2-44 Gene information for contactin family members Species Man

Mouse

Approved gene symbol CNTN1 CNTN2 CNTN3 CNTN4 CNTN5 CNTN6 Cntn1 Cntn2 Cntn3 Cntn4 Cntn5 Cntn6

Approved gene name Contactin 1 Contactin 2 (axonal) Contactin 3 (plasmacytoma associated) Contactin 4 Contactin 5 Contactin 6 Contactin 1 Contactin 2 Contactin 3 Contactin 4 Contactin 5 Contactin 6

Locus 12q11–q12 1q32.1 3p26

Database reference HGNC ID: 2171 HGNC ID: 2172 HGNC ID: 2173

3p26–25 11q21–22.2 3p26–p25 15 55.1 F 1 syntenic 6 43.5 6 E2 Unknown 6 E2

HGNC ID: 2174 HGNC ID: 2175 HGNC ID: 2176 MGI ID: 105980 MGI ID: 104518 MGI ID: 99534 MGI ID: 1095737 MGI ID: 3042287 MGI ID: 1858223

animals compared with younger animals. Hippocampal cells demonstrating an age‐related decrease in F3 expression include the pyramidal neurons of the CA1 region and the granule cells in the dentate gyrus (Shimazaki et al., 1998). F3 is not expressed in normal astrocytes but was recently demonstrated to be overexpressed in glioblastomas (Eckerich et al., 2005).

16.2.3 Isoforms and Protein Structure F3 has no intracellular domains but is attached to the membrane via a GPI anchor. The extracellular parts of both molecules consist of six Ig‐modules of the C2 type followed by four Fn3‐modules (> Figure 2-1) (Falk et al., 2002).

16.2.4 Posttranslational Modifications F3 contains sites for N‐linked glycosylation and expresses the HNK‐1 epitope (Kleene and Schachner, 2004).

16.2.5 Homophilic Interactions F3 does not engage in homophilic interaction (Falk et al., 2002) (> Table 2-45).

16.2.6 Heterophilic Binding Partners F3 interacts with the L1 family members L1, NrCAM, and neurofascin (Volkmer et al., 1998; Falk et al., 2002). F3 also binds tenascin‐C and ‐R, the transmembrane phosphatases RPTPa and b (Peles et al., 1995; Zeng et al., 1999), voltage‐gated sodium channel (Kazarinova‐Noyes et al., 2001; Liu et al., 2001; Shah et al., 2004), the neurexin family member Caspr (Falk et al., 2002), and the transmembrane protein Notch (Hu et al., 2003) (> Table 2-45).


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. Table 2-45 Binding partners for contactin family members CAM F3

TAG‐1

Extracellular binding partner Caspr L1 NrCAM Neurofascin RPTPa and b Notch TAG‐1 Tenascin‐C and –R Voltage‐gated sodium channels F3 Integrins L1 NCAM Neurocan Phosphacan TAG‐1 (trans)

Reference Falk et al. (2002) Falk et al. (2002) Morales et al. (1993); Volkmer et al. (1998) Volkmer et al. (1998) Peles et al. (1995); Zeng et al. (1999) Hu et al. (2003) Pavlou et al. (2002) Falk et al. (2002); Volkmer et al. (1998) Kazarinova‐Noyes et al. (2001); Liu et al. (2001); Shah et al. (2004) Pavlou et al. (2002) Felsenfeld et al. (1994) Malhotra et al. (1998) Milev et al. (1996) Milev et al. (1996) Milev et al. (1996) Furley et al. (1990); Alvarez‐Dolado et al. (2001); Freigang et al. (2000)

16.2.7 Signaling F3 has been reported to signal via the Notch receptor and triggers translocation of the intracellular domain of Notch to the nucleus (Hu et al., 2003).

16.2.8 Function F3 modulates axonal growth and guidance and mediates neuronal contact to glial cells, which influences axonal fasciculation. This is evident in F3‐knockout mice, which exhibit severe deficits in axonal projections in the cerebellum. Furthermore, F3‐knockout mice display a severe ataxic phenotype (defects in controlling voluntary movements), exhibit reduced growth, and only live around till P18 (Berglund et al., 1999; Falk et al., 2002). F3 has also been suggested to be important for memory formation by being important for long‐term depression (LTD) in the CA1 region of the hippocampus (Murai et al., 2002). (See also Sect. 18.3.8).

16.3 TAG‐1 16.3.1 Genes CNTN2, the gene encoding TAG‐1, has a 164‐bp region immediately upstream of the transcription initiation site, which is sufficient as a basal promoter, but additional regulatory regions are found further upstream of the initiation site. The 1.4‐kb upstream region contains various response elements (GRE, NRSE (neuron‐restrictive silencer element)) and putative binding sites for several transcription factors including AP‐2, Brn‐2, est‐1, NF‐kB, and Sp1 (Kozlov et al., 1995; Denaxa et al., 2003) (> Table 2-44).

16.3.2 Expression TAG‐1 is solely expressed in neurons. It is highly expressed in the cerebellum and spinal cord (Ogawa et al., 1996; Alvarez‐Dolado et al., 2001; Plagge et al., 2001). The protein is predominantly expressed during

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neural development. However, in the adult mouse it is expressed in low levels in cerebellum, olfactory bulb, and hippocampus (Wolfer et al., 1998).

16.3.3 Isoforms and Protein Structure TAG‐1 has no intracellular domains but is attached to the membrane via a GPI anchor. The extracellular parts of both molecules consist of six Ig‐modules of the C2 type followed by four Fn3‐modules (Falk et al., 2002).

16.3.4 Posttranslational Modifications TAG‐1 exhibits N‐linked glycosylation and contains carbohydrates with HNK‐1 epitopes.

16.3.5 Homophilic Interactions TAG‐1 is capable of forming homophilic binding (Furley et al., 1990; Felsenfeld et al., 1994). Homophilic trans‐interactions between TAG‐1 molecules can result in the cis‐interactions with L1, which subsequently leads to an interaction between L1 and the cytoskeletal protein ankyrin (Malhotra et al., 1998) (> Table 2-45).

16.3.6 Heterophilic Binding Partners TAG‐1 interacts with F3 and with the L1 family members L1, NrCAM, and neurofascin (Malhotra et al., 1998; Pavlou et al., 2002). TAG‐1 has been shown to interact with integrins (Felsenfeld et al., 1994) and the sulfate proteoglycans neurocan and phosphacan (Milev et al., 1996) (> Table 2-45). Furthermore, TAG‐1 is a putative sialic acid‐binding protein (Kleene and Schachner, 2004).

16.3.7 Signaling TAG‐1 has been reported to signal through the Src kinase Lyn, which in turn phosphorylates an unidentified 80‐kDa protein (Kasahara et al., 2000).

16.3.8 Function TAG‐1 promotes neuronal differentiation, neurite outgrowth, and regulates axonal guidance (Stoeckli et al., 1991; Buttiglione et al., 1998; Fitzli et al., 2000). Furthermore, studies have indicated that the molecule is important for the migration of cortical interneurons and neurons forming various precerebellar nuclei (Denaxa et al., 2001; Kyriakopoulou et al., 2002). However, results from a recent study utilizing TAG‐1‐deficient mice indicate that the protein is required for the survival of neurons of the precerebellar nuclei, but not for the migration of cortical interneurons (Denaxa et al., 2005). In the cerebellar cortex, TAG‐1 is under normal conditions expressed before F3. Transgenic mice, in which the TAG‐1 promoter regulates F3 expression, demonstrate alterations in cerebellar development caused by a reduction in the number of granule cells. Furthermore, these transgenic mice demonstrate several behavioral alterations including reduced exploratory activity, impaired motor activity, coordination, and learning. Hence, not only the expression of contactins, but also a correct spatiotemporal expression of the other proteins of the family is pivotal for proper development and function of the CNS (Bizzoca et al., 2003; Coluccia et al., 2004).


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16.4 Contactin‐3 Contactin‐3 was cloned in 1994. It is predominantly expressed in neurons including Purkinje cells of the cerebellum, granule cells of the dentate gyrus, and neurons in the superficial layers of the cerebral cortex (Connelly et al., 1994; Yoshihara et al., 1994). Plating cultured neurons on a coat of contactin‐3 promotes neurite outgrowth (Yoshihara et al., 1994).

16.5 Contactin‐4 Contactin‐4 was cloned in 1995. It is highly expressed in pyramidal cells in the CA1 region of the hippocampus (Yoshihara et al., 1995). In humans, it is also expressed in testis, uterus, thyroid, and small intestine (Hansford et al., 2003). In the mouse, it is highly expressed in the olfactory epithelium, where it appears around E14 and is expressed in high amounts between P(0) and P(7) (Saito et al., 1998). In humans, a splice variant of contactin‐4 (termed CNTN4A) has been identified. This isoform contains two Fn3‐modules attached to the membrane by a GPI‐anchor and is predominantly expressed in the cerebellum (Zeng et al., 2002). In the mouse, a soluble splice variant of contactin‐4 (termed BIG‐2A) has been identified. This isoform consists of six Ig‐modules and a single Fn3‐module, and lacks a GPI anchor. This form of contactin‐4 is predominantly expressed by mature sensory cells of the vomeronasal and olfactory neuroepithelium (Mimmack et al., 1997). Plating cultured neurons on a coat of contactin‐4 promotes neurite outgrowth (Yoshihara et al., 1995). Furthermore, disruption of contactin‐4 expression may be related to the rare gene disorder 3p deletion syndrome, which is characterized by developmental delay and growth retardation (Fernandez et al., 2004).

16.6 Contactin‐5 Contactin‐5 was cloned in 1996 (Ogawa et al., 1996). In humans, two isoforms have been identified, and the protein has been shown to be highly expressed in the amygdala and occipital lobe (Kamei et al., 2000). In the mouse and rat, the protein during adulthood is expressed in the central auditory pathways, and contactin‐5‐deficient mice demonstrate impaired neuronal activity in the auditory system (Ogawa et al., 2001; Li et al., 2003b).

16.7 Contactin‐6 Contactin‐6 was cloned from rats in 1996. It is expressed in the spinal cord, cerebrum, and cerebellum (Ogawa et al., 1996). In humans, the protein is predominantly expressed in the cerebellum (Kamei et al., 1998). In the mouse, the expression of the protein decreases in the cerebrum during development, whereas it increases in the cerebellum, where the protein is also expressed in the adult mouse. Mouse contactin‐6 has been reported to exist in two isoforms (Lee et al., 2000b). Contactin‐6‐deficient mice are viable and fertile, but demonstrate impaired motor coordination (Takeda et al., 2003).

17

The L1 Family

17.1 Introduction The L1 family of CAMs consists of four members: L1, NrCAM, neurofascin, and CHL1 (> Table 2-46). The proteins share a well‐conserved overall structure and are expressed from four genes believed to originate from one ancestral L1‐type gene as a result of sequential gene duplications (Pebusque et al., 1998).

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. Table 2-46 Nomenclature for L1 family members CAM name CHL1 L1 Neurofascin NrCAM

Protein and gene synonyms A530023M13Rik, CALL, L1CAM2 CAML1, CD171, HGNC:7086, HSAS, HSAS1, MASA, MIC5, S10, SPG1 D430023G06Rik, DKFZp686P2250, KIAA0756, mKIAA0756, NFASC hBravo, NgCAM‐related

L1 was originally showed to belong to the Ig superfamily by Moos and coworkers (1988). NrCAM was first identified as an adhesion molecule related to NgCAM (the chick homologue of L1) and was, on the basis of structural and sequential similarities with L1, classified as an L1 family member (Grumet et al., 1991). Neurofascin was cloned in 1992 (Volkmer et al., 1992), and the ‘‘close homologue of L1’’ (CHL1) was cloned in 1996 (Holm et al., 1996).

17.2 L1 17.2.1 Gene In humans, the L1 gene is located on chromosome X between the ALD and MeCP2 loci (Hlavin and Lemmon, 1991; Rosenthal et al., 1991, 1992). It consists of 29 exons and spans 13.9 kb (Kallunki et al., 1997). (See also > Table 2-47). . Table 2-47 Gene information for L1 family members Species Man

Approved gene symbol CHL1

NFASC NRCAM Chl1 L1cam

Approved gene name Cell adhesion molecule with homology to L1CAM (close homolog of L1) L1 cell adhesion molecule (hydrocephalus, stenosis of aqueduct of Sylvius 1, MASA (mental retardation, aphasia, shuffling gait, and adducted thumbs) syndrome, spastic paraplegia 1 Neurofascin Neuronal cell adhesion molecule Cell adhesion molecule with homology to L1CAM L1 cell adhesion molecule

Nfasc Nrcam

Neurofascin Neuron–glia–CAM–related cell adhesion molecule

L1CAM

Mouse

Locus 3p26


Xq28

HGNC ID: 6470

1q32.1 7q31 6 E1 X 29.51 A6‐B 1 70.0 12 22.0

GeneID: 23114 HGNC ID: 7994 MGI ID: 1098266 MGI ID: 96721 MGI ID: 104753 MGI ID: 104750

17.2.2 Expression L1 is mainly expressed in the nervous system but is also expressed in a developmentally regulated manner in a wide variety of tissues including the kidneys, immune system, urogenital tract, and digestive system (Thor et al., 1987; Kowitz et al., 1992; Kujat et al., 1995; Debiec et al., 1998). In the nervous system, one L1 isoform


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is specifically expressed on neuronal cells whereas another isoform is confined to nonneuronal cells such as Schwann cells, lymphocytes, and astrocytes. L1 expression is high along axonal pathways, and during development L1 is generally widespread throughout the PNS and CNS. In the adult, high L1 expression is found in the olfactory bulb, optic nerves, cerebellum, and hippocampus (Lyckman et al., 2000; Munakata et al., 2003).

17.2.3 Isoforms and Protein Structure The overall protein structure is conserved throughout the mammalian L1 family, which contains proteins consisting of six Ig‐modules of the C2‐type (denoted IgI–IgVI) followed by five Fn3‐modules (denoted Fn3I–Fn3V), a single transmembrane segment, and a cytoplasmic tail (Moos et al., 1988) (> Figure 2-1). Alternative splicing gives rise to two isoforms of L1. One isoform is exclusively expressed in neurons and contains a four‐amino acid sequence (RSLE), encoded by a small 12‐nucleotide exon, in the cytoplasmic tail (Miura et al., 1991). The other, nonneuronal, isoform is expressed in Schwann cells, astrocytes, melanocytes, and lymphocytes and lacks both the RSLE sequence in the cytoplasmic tail and a KGHHV sequence in the extracellular part (Takeda et al., 1996).

17.2.4 Posttranslational Modifications L1 can be phosphorylated at the cytoplasmic tail by the serine/threonine kinases p90rsk, CKII, Raf‐1, and ERK1/2. The p90rsk phosphorylation site is at S1152, the CKII phosphorylation site at S1181, C‐terminal of the RSLE sequence, and ERK1/2 can phosphorylate L1 at S1204 and S1248. The ERK1/2 phosphorylation sites are also found in NrCAM and neurofascin (Wong et al., 1996; Schaefer et al., 1999). L1 contains sites for N‐linked glycosylation and expresses the L2/HNK‐1, L3, L4, and L5 carbohydrate epitopes (see review by Krog and Bock, 1992).

17.2.5 Homophilic Interactions All members of the L1 family, except CHL1, share the ability to engage in homophilic binding (Hillenbrand et al., 1999) (> Table 2-48). L1 forms trans‐homophilic interactions in a manner involving the N‐terminal Ig‐modules (Miura et al., 1992). Deletion of the Ig2 module in L1 abolishes L1–L1 interactions and a 14‐amino acid sequence in this module has been identified as necessary for the interaction (Zhao and Siu, 1995; Zhao et al., 1998). It has also been speculated that L1 forms cis‐homodimers (reviewed by Kamiguchi and Lemmon, 1997) (> Table 2-48). Furthermore, it has been demonstrated that the first four Ig‐modules of L1 can adopt a horseshoe conformation. However, whether the molecule is extended or adopts a horseshoe conformation when it forms homo‐ and heterophilic interactions is unclear (> Figure 2-8); (see review by Haspel and Grumet, 2003).

17.2.6 Heterophilic Binding Partners Extracellular Heterophilic Binding Partners L1 forms cis‐interactions with several GPI‐linked receptors including TAG‐1, F3, and nectadrin (CD24) (Brummendorf et al., 1993; Kadmon et al., 1995; Malhotra et al., 1998; reviewed by Falk et al., 2002) (> Table 2-45). L1 also binds in cis to FGFR. The L1–FGFR interaction has been speculated to happen as a result of homophilic L1 trans‐binding, thus generating an L1–FGFR complex. The proposed model suggests an interaction between a CAM homology domain (CHD) in the FGFR and the Fn3IV module of L1, which results in the activation of FGFR (Doherty and Walsh, 1996; reviewed by Hall et al., 1996).

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. Table 2-48 Binding partners for L1 family members CAM L1

Extracellular binding partner L1 (cis and trans) CD9 CD24 (Nectadrin) F3 FGFR Integrin aIIbb3 Integrin aIIIb3 Integrin avb1 Integrin aVb3

Integrin a5b1

NrCAM

Neurofascin

CHL1 L1

NrCAM Neurofascin CHL1

Integrin a9b1 NCAM TAG‐1 Neuropilin‐1 SemaA3 NrCAM ALCAM F3 Neurofascin TAG‐1 Voltage‐gated sodium channels (b1 subunit) Neurofascin F3 NrCAM Syntenin‐1 TAG‐1 TN‐R Voltage‐gated sodium channels (b1 and b3 subunits) Integrin b1 (a1b1; a2b1) Actin Ankyrin AP‐2 Ankyrin SAP102 Ankyrin Ankyrin

Reference Miura et al. (1992); Zhao and Siu (1995); Zhao et al. (1998); Kamiguchi and Lemmon (1997) Schmidt et al. (1996) Kadmon et al. (1995) Falk et al. (2002) Doherty and Walsh (1996) Blaess et al. (1998) Felding‐Habermann et al. (1997) Felding‐Habermann et al. (1997) Montgomery et al. (1996); Felding‐Habermann et al. (1997); Blaess et al. (1998); Oleszewski et al. (1999); Silletti et al. (2000) Ruppert et al. (1995); Felding‐Habermann et al. (1997); Blaess et al. (1998); Oleszewski et al. (1999); Silletti et al. (2000) Silletti et al. (2000) Horstkorte et al. (1993) Malhotra et al. (1998) Castellani et al. (2002) Castellani et al. (2000) Mauro et al. (1992) DeBernado and Chang (1996) Morales et al. (1993) Volkmer et al. (1996) Suter et al. (1995) McEwen and Isom (2004) Hillenbrand et al. (1999) Volkmer et al. (1996) Volkmer et al. (1998) Koroll et al. (2001) Volkmer et al. (1998) Volkmer et al. (1998) Ratcliffe et al. (2001) Buhusi et al. (2003) Davis and Bennett (1994) Davis and Bennett (1994) Kamiguchi et al. (1998) Davis and Bennett (1994) Davey et al. (2005) Davis and Bennett (1994) Davis and Bennett (1994); Buhusi et al. (2003)


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. Figure 2-8 Extracellular L1 interactions. The many interactions mediated by L1 have been divided into ‘‘modular’’ and ‘‘cooperative’’ binding. Modular interactions involve a single L1‐module, and do not require assistance from other L1 modules. Cooperative interactions involve more than one L1‐module (the horseshoe formation involving Ig‐module 1–4) (Taken from Haspel and Grumet, 2003)

L1 binds in trans to NCAM. This interaction is believed to occur between oligomannosidic carbohydrates expressed by L1 and the IgIV module on NCAM (Horstkorte et al., 1993). The interaction requires homophilic NCAM binding, and subsequently strengthens homophilic L1 interactions (Kadmon et al., 1990a). L1 binds several integrins via RGD sequences located in IgIV; L1 interacts with integrin a5b1, avb1, avb3, aIIbb3, and aIIIb3 (Ruppert et al., 1995; Montgomery et al., 1996; Felding‐Habermann et al., 1997; Blaess et al., 1998; Oleszewski et al., 1999). Furthermore, the Fn3III‐module binds in an RGD‐independent manner to integrin avb3, a5b1, and a9b1 (Silletti et al., 2000). L1 has also been shown to interact with CD9 (a member of the tetraspanin family), neuropilin‐1, and the secreted semaphorin, semaA3 (Schmidt et al., 1996; Castellani et al., 2000, 2002; > Table 2-48). Finally, L1 has been shown to bind a2,3‐sialic acid in the Fn3I module (Kleene and Schachner, 2004), and the chicken orthologue of L1 has been shown to bind the ECM proteins laminin, phosphacan, and neurocan (see reviews by Hortsch, 1996; Haspel and Grumet, 2003). Intracellular Binding Partners L1 may be linked to the actin cytoskeleton through a C‐terminal binding site for ankyrin, but the protein may also interact directly with actin through an unknown linker protein, which binds to L1 close to the plasma membrane (Davis and Bennett, 1994; Dahlin‐Huppe et al., 1997; Wiencken‐ Barger et al., 2004). Furthermore, L1 binds the adaptor protein AP‐2, which is involved in endocytosis (Kamiguchi et al., 1998) (> Table 2-48).

17.2.7 Signaling Homo‐ and heterophilic binding by L1 family members may result in subsequent activation of intracellular signaling cascades. Homophilic L1 interactions may result in binding and activation of FGFR (Doherty and Walsh, 1996), subsequently leading to activation of PLCg and generation of IP3 and DAG from cleavage of PIP2. IP3 binds to Ca2þ channels in the intracellular calcium stores with a resulting increase

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in intracellular calcium concentration while DAG may activate PKC. DAG may also cause a rise in intracellular calcium concentration through binding to N‐ and L‐type calcium channels in the plasma membrane, which subsequently activates CAMII kinase; for reviews, see Kamiguchi and Lemmon (1997) and Crossin and Krushel (2000). L1 can activate Raf‐1 and p90rsk, thereby stimulating the MAPK pathway, which results in activation of transcription factors such as c‐Fos and CREB. In addition, PLD was recently demonstrated also to be activated by L1‐mediated activation of ERK (Watanabe et al., 2004), and the L1‐ mediated ERK activation also induces the expression of several gene products normally associated with motility and invasion (Silletti et al., 2004). ERK2 can phosphorylate L1 at S1204 and S1248. Hence, L1 signaling may regulate L1 function (Schaefer et al., 1999). The phosphorylation sites corresponding to S1204 and S1248 of L1 are also found in NrCAM and neurofascin, suggesting a similar signaling following activation of these proteins. The nonreceptor tyrosine kinase pp60c‐src has been suggested as a downstream mediator of L1 signaling, since L1‐mediated neurite outgrowth is inhibited in pp60c‐src‐deficient cells (Ignelzi Jr. et al., 1994).

17.2.8 Function In the nervous system, L1 is present on developing axons where it facilitates fasciculation by stimulating neurite extension along existing axons (Lemmon et al., 1989). During development L1, NrCAM, and neurofascin are involved in regulation of cell adhesion, neuronal migration, neurite outgrowth, growth cone morphology, axonal pathfinding, and LTP in the hippocampus (Kadmon et al., 1990a; Walsh and Doherty, 1997; for review see Hortsch, 1996). In both juvenile and adult animals, L1 has been implicated in regulation of synaptogenesis and synaptic plasticity (Saghatelyan et al., 2004). Furthermore, L1 and CHL1 promote neuronal survival. The effects can be observed when the extracellular parts of the proteins are used as substrates or are added in solution to cultures of cerebellar and hippocampal neurons. Hence, the neuroprotective effect is probably the result of trans‐interactions (Chen et al., 1999). L1‐deficient mice exhibit defects in axon guidance in the corticospinal tract and demonstrate abnormal development of the corpus callosum, reduced size of the hippocampus and cerebellum, and altered distribution of dopaminergic neurons (Cohen et al., 1998; Fransen et al., 1998a; Demyanenko et al., 1999, 2001). Mutations in the human L1 gene cause X‐linked hydrocephalus MASA syndrome (also referred to as the CRASH syndrome Fransen et al. (1998b)). CRASH syndrome patients are characterized by agenesis of the corpus callosum and mental retardation (Weller and Gartner, 2001).

17.3 NrCAM 17.3.1 Gene The NrCAM gene is located on chromosome 7 in humans at a locus believed to contain a tumor suppressor candidate gene (Lane et al., 1996). (See also > Table 2-47).

17.3.2 Expression Outside the CNS, NrCAM is expressed in several tissues including placenta, pancreas, and the adrenal tissues (Wang et al., 1998). In the nervous system, NrCAM is expressed in neural as well as nonneural cells such as Schwann cells. The protein is particularly highly expressed at the nodes of Ranvier (Custer et al., 2003).

17.3.3 Isoforms and Protein Structure NrCAM exists in a large number of isoforms produced by alternative splicing.


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17.3.4 Posttranslational Modifications NrCAM and neurofascin may be phosphorylated by tyrosine kinases (Garver et al., 1997; Custer et al., 2003).

17.3.5 Homophilic Interactions NrCAM is able to form homophilic interactions. The interactions have been reported to be dependent on divalent cations (Mauro et al., 1992) (> Table 2-48).

17.3.6 Heterophilic Binding Partners Extracellular Heterophilic Binding Partners NrCAM binds another member of the L1 family, neurofascin (Volkmer et al., 1996), and also binds the CAMs F3, TAG‐1, and ALCAM (Morales et al., 1993; Suter et al., 1995; DeBernado and Chang, 1996). Intracellular Binding Partners NrCAM binds ankyrin (Davis and Bennett, 1994) (> Table 2-48). Furthermore, the PDZ protein SAP102 was recently found to bind to the last three amino acids in the C‐terminal tail of NrCAM, and removal of these amino acids from NrCAM results in a protein with dominant negative effects on neurite outgrowth (Davey et al., 2005).

17.3.7 Function NrCAM stimulates neurite outgrowth. If the protein is used as a substrate, it stimulates neurite outgrowth through trans‐interactions with F3. In NrCAM‐expressing neurons, the protein stimulates neurite outgrowth through trans‐interactions with neurofascin (Volkmer et al., 1996). Differential display analysis has identified NrCAM as a protein expressed in neurons linked to reward and memory, and hence associated with substance abuse vulnerability. In rats, a single treatment with morphine leads to an upregulation of NrCAM expression. NrCAM‐deficient mice demonstrate a reduced preference for places where they can receive morphine, amphetamine, or cocaine compared with NrCAM‐expressing animals. Consistently, specific alleles of NRCAM are related to substance abuse in humans (Ishiguro et al., 2006).

17.4 Neurofascin 17.4.1 Gene In humans the neurofascin gene is located at chromosome 1q32 and spans 97.7 kb (Nagase et al., 1998). (See also > Table 2-47).

17.4.2 Expression In the nervous system, neurofascin is highly expressed at the nodes of Ranvier (Custer et al., 2003; Gollan et al., 2003) and is also located at unmyelinated axons in the PNS. At early stages of development, neurofascin is concentrated in developing fiber tracts (Rathjen et al., 1987; Hassel et al., 1997), whereas the expression is broader later in development (Moscoso and Sanes, 1995).

17.4.3 Isoforms and Protein Structure Neurofascin exists in a large number of isoforms produced by alternative splicing of NrCAM. As much as 50 different isoforms of neurofascin have been described, and their expression is regulated in a developmentally dependent manner (Hassel et al., 1997). The two main isoforms of neurofascin expressed in the

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nervous system are a 186‐kDa isoform, which is expressed on myelinated axons, and a 155‐kDa isoform, which lacks the Fn3III domain and mainly expressed in the PNS (Davis et al., 1996; Wang et al., 1998).

17.4.4 Posttranslational Modifications Extensive glycosylation of neurofascin has been reported with a proline‐, alanine‐ and threonine‐rich (PAT) region of 75 amino acids in close proximity of the plasma membrane, a specific target of O‐linked glycosylation (Volkmer et al., 1992).

17.4.5 Homophilic Interactions Neurofascin is able to form homophilic interactions (Hillenbrand et al., 1999) (> Table 2-48).

17.4.6 Heterophilic Binding Partners Extracellular Heterophilic Binding Partners Neurofascin interacts with NrCAM, and also binds the CAMs TAG‐1, F3, and the ECM glycoprotein TN‐R (Volkmer et al., 1998). The interaction with F3 can only take place when F3 is not interacting with Caspr (Gollan et al., 2003). The binding of neurofascin to TAG‐1 and TN‐R is regulated by the alternatively spliced PAT region, unique to neurofascin, suggesting that the heterophilic binding partners of neurofascin are differentially regulated with the expression of different neurofascin isoforms (Volkmer et al., 1998). Neurofascin has also been demonstrated to bind the protein syntenin‐1 (Koroll et al., 2001) Finally, the first Ig‐module and the second Fn3‐module of neurofascin binds the Ig‐module of the b1 and b3 subunits of voltage‐gated sodium channels at the nodes of Ranvier (Ratcliffe et al., 2001) (> Table 2-48). Intracellular Binding Partners Neurofascin binds ankyrin (Davis and Bennett, 1994) (> Table 2-48). Furthermore, doublecortin binds to the phosphotyrosine motif FIGQY (Kizhatil et al., 2002).

17.5 CHL1 17.5.1 Genes The human CHL1 gene is mapped at chromosome 3p26 and spans 212 kb. The predominant transcript is 8 kb (Wei et al., 1998) (See also > Table 2-47).

17.5.2 Expression CHL1 is expressed by neurons in the CNS and PNS as well as nonneural cells including Schwann cells, astrocytes, and oligodendrocytes. The expression pattern of CHL1 partially overlaps with that of L1 (Hillenbrand et al., 1999). Human CHL1 is highly expressed in the brain, heart, prostate, ovary, and intestines (Wei et al., 1998). Furthermore, it is expressed in some tumor cell lines including G361 melanoma cells and cervical carcinoma HeLa cells, whereas it not has been detected in leukemia, colorectal, or lung tumor cell lines (Wei et al., 1998). Rat CHL1 is widely expressed in all parts of the developing brain (Wei et al., 1998). The protein has been found to be overexpressed during axonal regeneration of motor neurons in vivo (Chaisuksunt et al., 2000; Zhang et al., 2000).


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17.5.3 Isoforms and Protein Structure The CHL1 exists in at least two isoforms, transmembrane isoform and soluble isoform. Thus, extracellular cleavage of the protein by the protease ADAM8 leads to soluble CHL1, which can stimulate neurite outgrowth in vitro (Hillenbrand et al., 1999; Naus et al., 2004). The protein contains several extracellular potential integrin‐binding sites (Wei et al., 1998).

17.5.4 Posttranslational Modifications CHL1 contains sites for O‐ and N‐linked glycosylations and expresses the HNK‐1 epitope (Holm et al., 1996; Hillenbrand et al., 1999).

17.5.5 Homophilic Interactions CHL1 does not form homophilic interactions (Hillenbrand et al., 1999).

17.5.6 Heterophilic Binding Partners Extracellular Heterophilic Binding Partners CHL1 is unique among L1 family proteins in that it does not engage in homophilic binding but exerts its adhesive effects solely through heterophilic interactions (Hillenbrand et al., 1999). Intracellular Binding Partners CHL1 binds ankyrin (Davis and Bennett, 1994) (> Table 2-48).

17.5.7 Signaling CHL1 has indirectly been shown to signal through the c‐Src, PI3, and MAP kinases. Thus, enhanced cell migrations mediated by CHL1–integrin interactions depend on the activation of these proteins (Buhusi et al., 2003).

17.5.8 Function Transwell assays have demonstrated that CHL1 potentiates integrin‐dependent haptotactic cell migration toward the ECM proteins collagen I, fibronectin, laminin, and vitronectin (Buhusi et al., 2003). Furthermore, CHL1 stimulates neuronal cell survival (see Sect. 19.2.8). A mutation in the signal peptide‐encoding sequence of CHL1 has been associated with schizophrenia, and a reduced expression level has been related to mental retardation (Sakurai et al., 2002; Frints et al., 2003; Chen et al., 2005). CHL1‐deficient mice are viable and fertile, but exhibit alterations in the hippocampus and olfactory system leading to various behavioral alterations. Furthermore, the lack of CHL1 leads to an upregulation of NCAM‐180 (Montag‐Sallaz et al., 2002; Frints et al., 2003; Pratte et al., 2003).

18

Down Syndrome Cell Adhesion Molecule and Down Syndrome Cell Adhesion Molecule‐Like 1

18.1 Introduction Down syndrome cell adhesion molecule (DSCAM) and the closely related Down syndrome cell adhesion molecule‐like 1 (DSCAML1) (> Table 2-49) are among the CAMs with the largest extracellular domains

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. Table 2-49 Nomenclature for DSCAM and DSCAML1 CAM name DSCAM DSCAML1

Protein and gene synonyms 4932410A21Rik, CHD2‐42, CHD2‐52 4921507G06Rik, 4930435C18Rik, KIAA1132, mKIAA1132

with their 16 Ig‐ and Fn3‐modules. DSCAM from mice and humans show 98% identity (Agarwala et al., 2001a), and human DSCAM and DSCAML1 show 64% identity extracellularly, and 45% identity in the cytoplasmic tail (Agarwala et al., 2001b). Extracellularly, but not intracellularly, the proteins also share strong homology with Drosophila DSCAM. Within the same species, DSCAM shares the closest homology to contactin‐4 (27% amino acid identity; 47% similarity (Yamakawa et al., 1998). Both DSCAM and DSCAML1 have been cloned independently by several groups. DSCAM was initially cloned from humans in 1998, and DSCAML1 was cloned a few years later (Yamakawa et al., 1998; Agarwala et al., 2001a, b) (> Table 2-50). . Table 2-50 Gene information for DSCAM and DSCAML1 Species Man

Mouse

Approved gene symbol DSCAM DSCAML1 Dscam Dscaml1

Approved gene name Down syndrome cell adhesion molecule Down syndrome cell adhesion molecule‐ like 1 Down syndrome cell adhesion molecule Down syndrome cell adhesion molecule‐ like 1

Locus 21q22.2–q22.3 11q22.2–q22.3

Database reference HGNC ID: 3039 HGNC ID: 14656

16 70.5 C 9B

MGI ID: 1196281 MGI ID: 2150309

18.2 Genes Both DSCAM and DSCAML1 are composed of at least 33 exons (Agarwala et al., 2001a, b, Barlow et al., 2002a), and both proteins exist in multiple isoforms as a result of alternative splicing of the original transcript (Yamakawa et al., 1998; Barlow et al., 2002a). Thus, DSCAML1 exists in at least three isoforms, depending on the inclusion of exon 3 or exon 4 (which encode the N‐ and C‐terminal halves of Ig‐module 2, respectively) (Barlow et al., 2002a). One isoform of DSCAM lacks the transmembrane region and contains a truncated cytoplasmic tail (Yamakawa et al., 1998). In humans, DSCAM mRNA transcripts 7.6, 8.5, and 9.7 kb have been identified, whereas the predominant transcript of DSCAML1 is 7.5 kb (Yamakawa et al., 1998; Barlow et al., 2002a). A 1.8 kb region upstream of DSCAM has been demonstrated to possess promoter activity and to drive the expression of a reporter gene in the CNS and other tissues (Barlow et al., 2002b).

18.3 Expression During mouse development, DSCAM is expressed in nonneuronal tissues such as liver, lungs, and limb buds. However, the protein is predominantly expressed in the nervous system. First, in the neuroepithelium and spinal cord and later, at E11.5, the expression is detectable throughout the brain. In adult brain, the protein is expressed in the spinal cord, cortex, mitral and granular layers of the olfactory bulb, CA1, CA2, and CA3 regions of the hippocampus, thalamus, and in Purkinje cell layer and deep nuclei of the cerebellum (Yamakawa et al., 1998; Agarwala et al., 2001a, c; Barlow et al., 2001b).


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In humans DSCAML1 is predominantly expressed in the brain during adulthood, but the protein is also found in other tissues. During mouse development, the protein is not detectable until around E17, where it becomes expressed in neuronal tissues. In the adult mouse, the protein is expressed in the dentate gyrus and in pyramidal cells of the hippocampus, in the olfactory bulb, and in the Purkinje cell layer of the cerebellum (Agarwala et al., 2001b). A comparison of the expression of DSCAM and DSCAML1 in the mouse has demonstrated that the two proteins exhibit inverse dorsal–ventral expression patterns in the developing spinal cord. Thus, DSCAM predominantly is expressed ventrally early in development, whereas DSCAML1 predominantly is expressed dorsally. Later in development, the two proteins are distributed evenly throughout the spinal cord. A similar phenomenon is found in the developing cortex, where the two proteins are expressed in different regions, and in the adult cortex DSCAM predominantly is expressed in the pyramidal neurons of layers three and five, whereas DSCAML1 predominantly is expressed in the granule cells of layer two (Barlow et al., 2002a). In primary cultures of cerebellar and hippocampal neurons, DSCAM is localized in both dendrites and axons (Agarwala et al., 2001c), whereas DSCAML1 in differentiated PC12 cells mainly is located in axons (Agarwala et al., 2001b).

18.4 Isoforms and Protein Structure Both DSCAM and DSCAML1 are composed of an extracellular domain consisting of ten Ig‐modules of the C2‐type and six Fn3‐modules, followed by a transmembrane region and a cytoplasmic tail. In addition to their length, the extracellular parts of the molecules are unusual in their organization. Thus, only the first nine Ig‐modules are membrane distal, whereas the last Ig‐module is located between Fn3‐module four and five (Yamakawa et al., 1998; Agarwala et al., 2001a). DSCAM contains around 2012–2013 amino acids, including a 397‐amino acid‐long cytoplasmic tail (Agarwala et al., 2001a). DSCAML1 contains 2053 amino acids, including a 441‐amino acid‐long cytoplasmic tail (Agarwala et al., 2001b). DSCAML1 contains one extracellular RGD sequence and a so‐called zinc‐binding region 2, a consensus sequence usually found in zinc carboxypeptidases (Agarwala et al., 2001b).

18.5 Posttranslational Modifications DSCAM and DSCAML1 contain 18 and 15 sites for potential N‐linked glycosylation, respectively (Agarwala et al., 2001a, b). The cytoplasmic tail of DSCAM and DSCAML1 contains several motifs including potential sites for phosphorylation and N‐myristoylation (Agarwala et al., 2001a, b; Barlow et al., 2001b).

18.6 Homophilic Interactions Both DSCAM and DSCAML1 have been demonstrated to be involved in trans‐homophilic interactions in a manner that mediates cellular adhesion and aggregation (Agarwala et al., 2000, 2001b) (> Table 2-51).

. Table 2-51 Binding partners for DSCAM and DSCAML1 CAM DSCAM DSCAML1 Dscam

Extracellular binding partner DSCAM (trans) DSCAML1 (trans) Pak1

Reference Agarwala et al. (2000) Agarwala et al. (2001b) Li and Guan (2004)

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18.7 Heterophilic Binding Partners 18.7.1 Intracellular Binding Partners Human DSCAM has been demonstrated to interact with the serine/threonine kinase Pak1 (> Table 2-51). The interaction takes place between amino acid residues 1637–1791 of DSCAM and residues 351–400 within the kinase domain of Pak1. Binding of the GTPase Rac to the CRIB (Cdc42/Rac1 interactive binding)‐domain of Pak1 strengthens the DSCAM–Pak1 interaction (Li and Guan, 2004).

18.8 Signaling The interaction between DSCAM and Pak1 leads to the phosphorylation of Pak1 at S199/204 and subsequent alterations in cell shape and activation of MAPKs, JNK and p38 (> Figure 2-9) (Li and Guan, 2004).

18.9 Functions DSCAM and DSCAML1 are likely to be involved in the guidance and migration of cells during development. Thus, both proteins are expressed by neuroepithelial cells and are believed to be expressed by migrating neural crest cells (Yamakawa et al., 1998; Barlow et al., 2002a). In humans, DSCAM is located on chromosome 21 in the region critical for some of the neuropathological phenotypes related to Down syndrome. Consistently, the protein has been shown to be overexpressed in brains of people with Down syndrome. In Purkinje cells, the overexpression is age independent, whereas an overexpression in cortical neurons predominantly is found during adulthood. In people with Down syndrome an dementia, the protein has been found in senile plaques (Saito et al., 2000). Furthermore, the gene has been proposed also to be a candidate gene for Down syndrome congenital heart disease (DS‐CHD) (Barlow et al., 2001a).

19

Sidekick 1 and 2

19.1 Introduction Sdk1 and Sdk2 (Sidekick 1 and 2) (> Table 2-52) were cloned from humans and mice in 2002. They are orthologues of Drosophila sdk, with which they share 35% identity at the protein level (Yamagata et al., 2002) (> Table 2-53).

19.2 Expression No general analyses of the expression of Sdks have been published. However, their expression has been studied in detail in the inner plexiform layer of the retina, where dendrites of retinal ganglion cells are localized. Here, Sdks are located in synapses, whereas they are absent from the somata of these neurons (Yamagata et al., 2002). Sdk1 and Sdk2 have also been shown to be expressed in the kidneys. During mouse development they are initially expressed in the uretic buds and bud‐derived tissues, whereas they in the adult mouse are expressed in the glomeruli (Kaufman et al., 2004).

19.3 Isoforms and Protein Structure Sdk1 and Sdk2 have the same general structure and are 59% identical. They consist of six Ig‐modules of the C2‐type, 13 Fn3‐modules, a single transmembrane region, and a 200‐amino acid‐long cytoplasmic tail.


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. Figure 2-9 DSCAM Signaling. DSCAM binds directly to Pak1. Pak1 can interact with the small GTPases, Rac and Cdc42, through its CRIB domain. Furthermore, DSCAM may form homophilic cis‐interactions. The various protein interactions lead to activation of the MAPK, JNK. CRIB, Cdc42/Rac interaction/binding motif; DID, DSCAM‐ interacting domain; PID, Pak‐interacting domain (Modified from Li and Guan, 2004)

. Table 2-52 Nomenclature for SDK1 and SDK2 CAM name Sidekick 1 Sidekick 2

Protein and gene synonyms FLJ31425, 6720466O15Rik FLJ10832, KIAA1514, 4632412F08Rik, 5330435L01Rik

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. Table 2-53 Gene information for SDK1 and SDK2 Species Man Mouse

Approved gene symbol SDK1 SDK2 Sdk1 Sdk2

Approved gene name Sidekick homolog 1 (chicken) Sidekick homolog 2 (chicken) Sidekick homolog 1 (chicken) Sidekick homolog 2 (chicken)

Locus 7p22.3 17q21.31 5 syntenic 11 syntenic

Database reference HGNC ID: 19307 HGNC ID: 19308 MGI ID: 2444413 MGI ID: 2443847

The C‐terminal end of both proteins contains a conserved sequence, GFSSFV, which is expected to bind PDZ domain‐containing proteins. Furthermore, both Sdks contain a potential carbohydrate‐binding lectin homology sequence in their most membrane‐proximal Fn3‐modules (Yamagata et al., 2002).

19.4 Homophilic Interactions Sdk1 and Sdk2 are involved in homophilic trans‐interactions, but do not appear to form heterophilic Sdk1– Sdk2 interactions (Yamagata et al., 2002) (> Table 2-54). Recently, it was demonstrated that the first two . Table 2-54 Binding partners for SDK1 and SDK2 CAM SDK1 SDK2

Extracellular binding partner SDK1 (trans) SDK2 (trans)

Reference Yamagata et al. (2002); Hayashi et al. (2005) Yamagata et al. (2002); Hayashi et al. (2005)

Ig‐modules of Sdk1 and Sdk2 are sufficient and necessary for the formation of homophilic trans‐ interactions, which appear to be mediated by reciprocal interactions between the Ig1 and the Ig2 modules (Hayashi et al., 2005). No heterophilic interactions involving Sdk1 or Sdk2 have been published.

19.5 Function The study of Sdk1 and Sdk2 expression in the inner plexiform layer of the retina and surrounding regions has demonstrated that these proteins are involved in laminar‐specific synapse formation. In the retinal ganglion cell layer, the two proteins are found in nonoverlapping subsets of neurons, and in the inner plexiform layer (which can be divided into five zones, S1–5), Sdk1 is predominantly found in layer S4, whereas Sdk2 is predominantly found in layer S2. It is believed that the two proteins mediate specific interactions between pre‐ and postsynaptic terminals of Sdk‐positive cells (Yamagata et al., 2002; reviewed by Abbas, 2003). In addition, Sdk1 is upregulated in the kidneys of transgenic mice expressing the HIV‐1 genome and has therefore been related to HIV‐associated nephropathy (HIVAN) (Kaufman et al., 2004).

20

Putative Neural Adhesion Molecules

20.1 IGSF9 IGSF9 (immunoglobulin superfamily member 9) was cloned from humans and mice in 2002 (> Table 2-55 and > 2-56). Both genes consist of 21 exons, which pairwise are at least 75% homologous, and the predicted


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. Table 2-55 Nomenclature for putative neural adhesion molecules CAM name IGSF9 NOPE PUNC TMEM25

Protein and gene synonyms KIAA1355, Nrt1, Dasm1, dendrite arborization and synapse maturation 1 9330155G14Rik, mKIAA1628 FLJ14399, 0610039J01Rik

. Table 2-56 Gene information for putative neural adhesion molecules Species Man

Approved gene symbol IGSF9

Mouse Man Mouse Man

Igsf9 NOPE Nope PUNC

Mouse Man Mouse

Punc TMEM25 Tmem25

Approved gene name Immunoglobulin superfamily, member 9 Neighbor of Punc E11 Putative neuronal cell adhesion molecule Transmembrane protein 25

Locus 1q22–q23


1 94.0 15q22.31 9 syntenic 15q22.3–q23

MGI ID: 2135283 Locus ID: 57722 MGI ID: 1858497 HGNC ID: 9700

9 40.0 D‐E1 11q23.3 9 syntenic

MGI ID: 11202390 HGNC ID: 25890 MGI ID: 1918937

1179‐amino acid‐long proteins exhibit 87% identity (91% similarity). The structure of the protein resembles NCAM, consisting extracellularly of five Ig‐modules and two Fn3‐modules, followed by a transmembrane region and a cytoplasmic tail (amino acids 760–1179) (Doudney et al., 2002). The C terminus of the cytoplasmic tail contains a type I PDZ‐module‐binding motif (‐TLL) (Shi et al., 2004a). IGSF9 shares homology with a number of proteins. The highest homology exists between IGSF9 and the Drosophila melanogaster protein Turtle (31% identity and 50% similarity for the EC modules), but the protein also shares homology to hemicentin (41% similarity), fibulin‐6 (41% similarity), NCAM (41% similarity), neogenin (40% similarity), and NrCAM (37% similarity) (Doudney et al., 2002). In humans, transcription of the protein has been observed in various fetal tissues both at week 8 and week 14. In mouse, three mRNA transcripts for the protein have been identified, and of these, the longest is brain specific. The protein is expressed from E7.5, and transcription is intense within the dorsal root ganglia, trigeminal ganglia, and olfactory epithelium. At E18 the protein is highly expressed in the brain, including the hippocampus, cerebral cortex, and cerebellum. (Doudney et al., 2002; Shi et al., 2004a). Immunostaining indicates that the protein has a punctate distribution in cell bodies and dendrites of cortical and hippocampal neurons and cerebellar Purkinje cells (Shi et al., 2004a). Along dendrites the expression of IGSF9 partially overlaps with that of the subunits GluR2 and GluR4 of the AMPA receptor (Shi et al., 2004b). IGSF9 has been shown to interact with the two synaptic PDZ module‐containing proteins Shank and S‐SCAM, whereas it does not coimmunoprecipitate with PSD‐95, GRIP, SAP102, or SAP97 (Shi et al., 2004b). No extracellular homophilic or heterophilic binding partners for IGSF9 have been demonstrated. Knockdown of IGSF9 and deletion of the PDZ‐binding C‐terminal region reduce dendrite—but not axon—outgrowth, and impair AMPA‐R‐mediated—but not NMDA‐R‐mediated—synaptic transmission (Shi et al., 2004a, b). For a review of Igsf9, see Falls (2005).

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20.2 Punc Punc (putative neuronal cell adhesion molecule) was identified in mice and humans in 1998 and 1999, respectively, and the orthologues are 72% similar and 80% identical (> Table 2-55 and > 2-56). The protein belongs to the IgSF. It contains four Ig‐modules and two membrane‐proximal Fn3 modules, a transmembrane region, and a cytoplasmic tail. Punc exhibits homology to the guidance molecule DCC. Thus, the second, third, and fourth Ig‐module of Punc is 41, 42, and 47% identical to the respective modules of DCC (Salbaum, 1998, 1999). In addition, Punc exhibits homology to Nope (see below) (Salbaum and Kappen, 2000). The gene consists of 15 exons, of which exon 13 encodes the predicted transmembrane region, and exons 14 and 15 encode the cytoplasmic tail. The protein seems to exist in two isoforms, 793 and 813 amino acids, respectively, depending on whether the 60 bp‐long exon 12 is excluded or included in the transcript. Interestingly, this exon encodes the only putative site for N‐linked glycosylation, a sequence which is conserved between humans and mice (Salbaum, 1998, 1999). In the mouse, the protein is expressed in the neuroectoderm, and later, in the lateral plate mesoderm. In the neural tube, the level of expression appears to be correlated with periods of early cell proliferation (Salbaum, 1999). By E15.5, and in the adult, the expression is largely confined to the brain and inner ear. The strongest expression appears to be in the so‐called Bergmann glia in the cerebellar cortex (Yang et al., 2001a). Experiments with Punc‐null mice have demonstrated that the protein is not required for normal embryogenesis. Punc‐deficient animals are viable, fertile, and have—despite the lack of Punc in the inner ear—an apparently normal hearing. However, the animals demonstrate reduced motor coordination (identified as a significant reduction in the retention time on the so‐called Rota‐rod) (Yang et al., 2001a). Punc has not yet been demonstrated to be involved in any homo‐ or heterophilic interactions.

20.3 Nope Nope (neighbor of Punc E11) was cloned in the mouse in 2000 (> Table 2-55 and > 2-56). It is a 1252‐ amino acid‐long IgSF protein consisting of four Ig‐modules followed by five membrane‐proximal Fn3 modules, a single transmembrane region, and a 274‐amino acid‐long cytoplasmic tail. From the second Ig‐module to the first Fn3‐module Nope shares 45% homology to Punc, and the protein also exhibits homology with neogenin and DCC. The expression of Nope in the mouse can be detected from E9.5 and is expressed in all skeletal muscles. From E15.5 the protein is also expressed in the nervous system, predominantly in the ventricular zone of the brain and in the ventral midline of the diencephalon. In the adult brain the protein is expressed in the hippocampus, piriform cortex (a part of the olfactory system), thalamus, and cerebellum. Nope has not yet been demonstrated to be involved in any homo‐ or heterophilic interactions.

20.4 TMEM25 TMEM25 (transmembrane protein 25) was cloned from humans and mice in 2004 (> Tables 2-55 and > 2-56). In these two species the protein exhibits 91% identity. TMEM25 exists in at least two isoforms obtained by alternative splicing, a 366‐amino acid‐long transmembrane protein and a 322‐amino acid‐long secreted isoform. Codon 42–112 encodes a putative C‐2 type Ig‐module. TMEM25 is expressed in the brain including cerebellar cortex and hippocampus as well as in neuroblastoma, brain tumors, and gastric cancer (Katoh and Katoh, 2004). TMEM25 has not yet been demonstrated to be involved in any homo‐ or heterophilic interactions.


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Neurosecretory Protein Trafficking and Dense‐Core Granule Biogenesis in Neuroendocrine Cells

T. Kim . M. Gondre´‐Lewis . I. Arnaoutova . N. Cawley . Y. Peng Loh

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154

2 Sorting of Neuropeptides and BDNF to the RSP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 2.1 Sorting Signals in Neuropeptides and BDNF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155 2.2 Carboxypeptidase E: A Sorting Receptor for Neuropeptides and BDNF . . . . . . . . . . . . . . . . . . . . . . . . . 157 3

Trafficking of Proneuropeptide‐Processing Proteases to Secretory Granules . . . . . . . . . . . . . . . . . . . 157

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Secretory Granule Biogenesis in Neuroendocrine Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Role of Cholesterol in Secretory Granule Biogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Granulogenic Proteins: Assembly Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 Proteins Regulating DCG Biogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164

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Neurosecretory protein trafficking and dense‐core granule biogenesis in neuroendocrine cells

Abstract: Neuropeptides and brain‐derived neurotrophic factor (BDNF) are secreted from neurons in an activity‐dependent manner through the regulated secretory pathway (RSP). Neuropeptides and BDNF, initially synthesized as a proform, are sorted at the trans‐Golgi network (TGN) along with their processing enzymes, prohormone convertases (PC) 1/3 and 2 and carboxypeptidase E (CPE), into the dense‐core secretory granules (DCGs) for secretion. Consensus RSP sorting signals have been identified in proopiomelanocortin (POMC), insulin, and BDNF, which are sufficient and necessary for targeting these proteins to secretory granules. These signals are conformation‐dependent and consist of a pair of acidic amino acids 10–15 A˚ apart and an aliphatic hydrophobic amino acid 5–6 A˚ away from each of the acidic residues. The acidic residues in the motif interact with a sorting receptor, membrane CPE, at the TGN to effect sorting into granules. CPE‐deficient mouse models verify the functioning of the CPE‐mediated sorting mechanism in vivo. Other less well studied putative proneuropeptide sorting motifs and mechanisms are also discussed. The processing enzymes, however, are sorted to the DCGs by insertion of their C‐terminal domain into cholesterol–sphingolipid‐rich membrane microdomains (lipid rafts), which are the proposed sites of budding to form the DCGs at the TGN. In addition to sorting of cargo to the DCGs, molecules driving the budding of the DCGs from the TGN and regulating granule quantity are important components for regulated secretion of neuropeptides. The granins, major proteins in DCGs, as well as proneuropeptides form aggregates that appear to be able to provide the necessary driving force to form DCGs. In addition, cholesterol plays a critical role in the pinching off of the granules at the TGN, as evidenced from cholesterol depletion studies in an endocrine cell line and in cholesterol‐deficient mouse models. Finally, studies have identified chromogranin A (CgA) as a regulator of quantitative DCG biogenesis in neuroendocrine cells. CgA acts by regulating the stability of secretory granule proteins in the Golgi apparatus, thereby controlling the number of DCGs that can be formed. List of Abbreviations: BDNF, brain-derived neurotrophic factor; RSP, regulated secretary pathway; TGN, trans-Golgi network; PC, prohormone convertases; CPE, carboxypeptidase E; DCGs, dense-core secretory granules; POMC, proopiomelanocortin; CgA, chromogranin A; ISGs, immature secretory granules; NMR, nuclear magnetic resonance; NGF, nerve growth factor; SLOS, Smith–Lemli–Opitz syndrome; Sc5d, lathosterol desaturase; GUVs, giant unilamellar vesicles; CgB, chromogranin B; SecII, secretogranin II; PTB, Polypyrimidine-tract binding protein; ICA512/IA2, receptor-tyrosine-phosphatase-like protein

1

Introduction

Neurons secrete classical neurotransmitters, neurotrophins (NT3, BDNF), and neuropeptides in an activity‐dependent manner through a regulated secretory pathway (RSP) for various synaptic functions. The secretion of these molecules depends on the formation of small clear vesicles or dense‐core secretory granules (DCGs). Therefore the regulation of secretory granule biogenesis is critical for neuronal function. In addition, the sorting and packaging of cargo into these granules for secretion represents another critical step in bringing about activity‐dependent secretion of neuropeptides and brain‐derived neurotrophic factor (BDNF) for synaptic activity. Neuropeptides and BDNF are synthesized as larger precursors at the rough endoplasmic reticulum and trafficked to the trans‐Golgi network (TGN) where they are sorted and selectively packaged along with the processing enzymes into immature granules of the RSP. These precursors are processed to bioactive neuropeptides and mature BDNF, respectively, in the immature secretory granules (ISGs), stored and secreted upon stimulation. Processing occurs at dibasic cleavage sites and is carried out by a family of processing enzymes known as prohormone convertases (PCs). Over the last 8 years, a mechanism for sorting neuropeptide and BDNF precursors has been elucidated and much is also known about the trafficking of the processing enzymes into ISGs. However, the understanding of the regulation of formation of granules is only just emerging. Granins have long been proposed to be ‘‘granulogenic.’’ Studies have indicated that the granins and other prohormones act as assembly factors in DCG biogenesis. More recently, chromogranin A (CgA), present in all neuroendocrine and endocrine cells with the exception


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of somatotrophs and lactotrophs, has been shown to be an important regulatory factor, in addition to its granulogenic role in DCG formation. In this chapter, we review the mechanisms of sorting and packaging of neuropeptides, BDNF, and their processing enzymes, into granules of the RSP. We also discuss current knowledge of the factors governing the physical formation of DCGs and the regulation of DCG biogenesis.

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Sorting of Neuropeptides and BDNF to the RSP

2.1 Sorting Signals in Neuropeptides and BDNF Neuropeptide and BDNF precursors are sorted at the TGN into ISGs. Two hypotheses have been proposed for the mechanism of sorting neuropeptides into the ISGs. The ‘‘passive’’ sorting mechanism proposes that the neuropeptide precursors uniquely aggregate at the TGN under conditions of low pH and high Ca2þ and are therefore segregated from other non‐RSP proteins. The aggregate then binds to the TGN membrane either through nonspecific protein–protein interaction or through protein–lipid interaction and is then packaged into the ISGs (Tooze, 1991; Arvan and Castle, 1992). Evidence for this hypothesis comes from in vitro studies showing aggregation of various prohormone substrates at low pH and high Ca2þ and demonstration that these aggregates associate with membranes (Rindler, 1998; Dannies, 1999). Such a sorting mechanism does not require any specific sorting signal or receptor. The other is the ‘‘active’’ sorting mechanism, which proposes that a specific sorting signal exists on proneuropeptides and BDNF that interacts with a specific receptor, either a protein or a lipid at the TGN. Then budding occurs at the site of interaction at the TGN membrane to form the ISGs. This is known as ‘‘sorting for entry,’’ which occurs in neurons and neuroendocrine cells. The other ‘‘active’’ sorting mechanism is ‘‘sorting by retention,’’ which has been demonstrated so far only in endocrine cells. Proinsulin for example is sorted by retention from exit through the constitutive‐like pathway in the ISGs by binding of the sorting signal to a sorting receptor in pancreatic b‐cells (Dhanvantari et al., 2003). Several studies have identified RSP sorting signals in proteins destined for this pathway. This includes proneuropeptides: proopiomelanocortin (POMC), proinsulin; and BDNF (Cool et al., 1995; Zhou et al., 1999; Dhanvantari et al., 2003; Lou et al., 2005). > Figure 3-1a–c shows the sorting motif identified in the X‐ray crystal structure of insulin and BDNF and nuclear magnetic resonance (NMR) structure for POMC. The motif consists of a three‐dimensional conformational structure, with two acidic and two hydrophobic residues. The side chains of the two acidic residues are always exposed on the surface and the molecular distances between the four residues are very similar for the motif in each of the molecules illustrated (> Figure 3-1d ). The molecular distance between the acidic residues ranges between 9 and 16 A˚ and distances of the hydrophobic residues range between 5 and 8 A˚ from the acidic residues. In the case of proinsulin, hexamerization resulted in the presentation of an alternative but similar motif contributed by the same residues on the ‘‘A’’ chain of two adjacent dimers (see Dhanvantari et al., 2003). Site‐directed mutagenesis studies on POMC (Loh et al., 2004), proinsulin (Dhanvantari et al., 2003), and BDNF (Lou et al., 2005) have shown that the four residues in the motif are necessary for correct sorting of these molecules to the RSP. Mutation of the residues in the motif led to constitutive secretion of the mutant prohormone and immunocytochemical analyses showed little or no localization in punctate granules, indicating missorting when the motif was mutated. The stringency of the motif was demonstrated by the lack of one of the acidic residues in nerve growth factor (NGF) that would otherwise contain a complete motif similar to BDNF (> Figure 3-1e). NGF is largely secreted constitutively, but when the hydrophobic residue Val20 was substituted for an acidic residue, Glu20, to form the motif, it was targeted to the RSP (Lou et al., 2005), showing the specificity of the motif. This type of motif has also been found by molecular modeling to be present in other proneuropeptides, e.g., provasopressin and proenkephalin, but they have not been studied experimentally in detail. A search for a similar motif in albumin, a constitutively secreted protein, did not reveal such a motif in this molecule, indicating specificity for regulated secretory proteins.

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. Figure 3-1 Structural analysis of the sorting signals of three prohormones/proneuropeptides trafficked to the regulated secretory pathway. X‐ray crystal structures of (a) insulin and (b) brain‐derived neurotrophic factor (BDNF) and (c) nuclear magnetic resonance (NMR) structure of N‐POMC1–26 sorting signal domains, consisting of two acidic residues and two aliphatic hydrophobic residues. A comparison of the distances between each residue is shown in (d), where distances are in Angstroms. The top numbers are for proinsulin, the middle numbers are for BDNF, and the bottom numbers are for N‐POMC1–26. (e) The X‐ray crystal structure of NGF, which is constitutively secreted from cells, is very similar to that of BDNF; however, a complete sorting signal is not present due to the absence of a critical acidic residue in the Val20 position. When mutated to a Glu20, the mutant NGF(V20E) was sorted to the RSP (Lou et al., 2005)

While our studies have identified a motif in the mature BDNF domain of proBDNF that is required for sorting to the RSP (Lou et al., 2005), others have suggested that the prodomain contains information important for targeting proBDNF to the RSP for activity‐dependent secretion (Egan et al., 2003; Chen et al., 2004). These studies showed that a single nucleotide polymorphism in the BDNF gene leading to a valine‐ to‐methionine substitution at codon 66 in the prodomain (proBDNFmet) resulted in decreased activity‐ dependent secretion of BDNF. Immunohistochemical localization and secretion studies revealed significant accumulation of proBDNFmet in the Golgi apparatus and no elevated constitutive secretion when transfected into hippocampal and cortical neurons (Egan et al., 2003). However, little is known about the effect of the valine‐to‐methionine substitution on the structure of the prodomain that could provide a clue to the mechanism for the inefficient sorting of proBDNFmet to the RSP. On the basis of theoretical analysis of the amino acids and secondary structures of 15 proteins sorted to the RSP, Kizer and Tropsha (1991) proposed that a putative motif may consist of two or more leucines on


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one side of a highly amphipathic a‐helix, with a serine or threonine positioned N‐terminal to the leucines. Indeed, a sorting signal for prosomatostatin composed of an amphipathic a‐helix with two critical leucines has been demonstrated (Mouchantaf et al., 2001). Other studies have reported that structural features such as dibasic residues in prorenin (Brechler et al., 1996; Feliciangeli et al., 2001; Feliciangeli and Kitabgi, 2002) may play a role in directing sorting to the RSP. Progastrin was also shown to depend on two domains for sorting to the RSP: its dibasic cleavage site as the primary regulator and an acidic domain that appears to work synergistically with the cleavage site domain (Bundgaard et al., 2004).

2.2 Carboxypeptidase E: A Sorting Receptor for Neuropeptides and BDNF A search for a sorting receptor began with the use of the POMC sorting signal (POMC1–26) as a ligand in an affinity column and bovine pituitary secretory granule membranes as a source of a putative receptor. By such a procedure, the membrane form of carboxypeptidase E (CPE) (> Figure 3-2a) was identified as a specific binding protein for the POMC sorting signal (Cool et al., 1997). CPE has been shown to be a transmembrane protein associated with detergent‐resistant cholesterol–sphingolipid‐rich microdomains (lipid rafts) at the TGN (Dhanvantari et al., 2002). Molecular modeling of CPE (> Figure 3-2b) revealed an exposed domain, unique to CPE, between residues Asn247 and Thr272 that contained two basic amino acids, Arg255 and Lys260, whose side chains were exposed on the surface of the molecule (Dhanvantari et al., 2002). Molecular docking studies showed that the distance between the side chains of these two basic residues was appropriate for interaction with the two acidic residues in the sorting signal of POMC, proinsulin, and BDNF (> Figure 3-2c–e). Binding studies with N‐POMC1–26, which contain the sorting signal, showed highly specific binding of this peptide with recombinant CPE expressed in insect cells (Zhang et al., 1999). Substitution of amino acids DLEL in the POMC motif for ASAS in the N‐POMC1–26 peptide eliminated the binding. Furthermore, expression of a mutant CPE with Arg255 and/or Lys260 mutated to Ala in insect cells also eliminated binding of N‐POMC1–26 to these cell membranes (Zhang et al., 1999). Comparison of the binding of BDNF and NGF to recombinant CPE showed specific binding of BDNF but not NGF that lacks the complete sorting motif (Lou et al., 2005). Transfection of a dominant negative CPE (247–272)‐GFP fusion protein into INS‐1 cells, a pancreatic b‐cell line, diminished sorting of proinsulin to the RSP, compared with GFP alone, indicating interaction of the CPE domain with the proinsulin sorting signal ex vivo (Dhanvantari et al., 2003). Finally, POMC sorting to the RSP was eliminated in the pituitary of a mutant mouse, Cpefat/fat, that lacked CPE due to degradation in the pituitary (Cool et al., 1997; Shen and Loh, 1997). On the other hand, proinsulin sorting to the RSP was unaffected in these mice (Irminger et al., 1997), which showed significant amounts of mutant proCPE in the pancreatic b‐cell cells, in contrast to the pituitary (Naggert et al., 1995). Recently a CPE knockout mouse was generated (Cawley et al., 2004) and BDNF was shown to be missorted to the constitutive pathway in hippocampal and cortical neurons of these mice (Lou et al., 2005). All these studies demonstrate that CPE acts as a sorting receptor for targeting a number of proneuropeptides and proBDNF to the RSP (see > Figure 3-3).

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Trafficking of Proneuropeptide‐Processing Proteases to Secretory Granules

The proteolytic enzymes involved in processing of proneuropeptides to their bioactive peptides are the proprotein convertases (PCs), of which the major ones are PC1/3 and PC2. These enzymes cleave at paired basic residues (Thomas et al., 1991). Following the action of these proteases, CPE cleaves off the basic residues at the C terminus of the peptides liberated from the precursor (Fricker, 1988). PC1/3, PC2, and CPE exist as soluble proteins, but a significant population have also been shown to have a transmembrane orientation, with a cytoplasmic tail (> Figure 3-4) and are associated with lipid rafts at the TGN (Dhanvantari et al., 2002; Arnaoutova et al., 2003a; Assadi et al., 2004). Trafficking of PC1/3, PC2, and CPE have been extensively studied. Deletion mutagenesis studies have identified three domains at the C terminus of PC1/3 that are sufficient for sorting of this enzyme to the RSP

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. Figure 3-2 Structural analysis and molecular interaction of carboxypeptidase E with the sorting signals of POMC, proinsulin, and brain‐derived neurotrophic factor (BDNF). CPE is synthesized as (a) prepro‐CPE of 476 amino acids. The signal peptide (SP) is removed in the endoplasmic reticulum and the 14‐amino‐acid proregion (PRO) is removed in the granules of the regulated secretory pathway (RSP). The last 25 amino acids of CPE (TM) confer membrane association of CPE in both a peripheral and an intrinsic transmembrane manner. Modeling of (b) CPE based on the crystal structure of the enzymatically active domain of carboxypeptidase D reveals the surface availability of the sorting signal binding site, which involves two basic residues, Arg255 and Lys260. Molecular docking of the sorting signals of (c–e) POMC, proinsulin, and BDNF to the sorting signal binding site of CPE demonstrate the complimentarity between the two acidic residues of the sorting signals in each case and the two basic residues of the sorting signal binding site on CPE

(Zhou et al., 1995; Lusson et al., 1997; Rovere et al., 1999). These regions include the transmembrane domain (residues 616–632) (Arnaoutova et al., 2003a) and residues 667–713 and 711–753, which reside in the cytoplasmic tail (Jutras et al., 2000). Of these segments, only the transmembrane domain was found to be sufficient and necessary for sorting PC1/3 to the secretory granules within the context of the whole


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. Figure 3-3 Model for the involvement of CPE in the trafficking of prohormones to the regulated secretory pathway (RSP). CPE becomes a raft‐associated protein in the late Golgi apparatus through interaction of its C terminus with the membrane. The acidic environment and elevated calcium concentration of the trans‐Golgi network (TGN) initiates the loose aggregation of luminal prohormones, which bind to the membrane‐anchored CPE via the sorting signals. As the vesicle buds at sites where rafts have coalesced, CPE carries the aggregated cargo into the immature granule where processing begins followed by condensation of the peptide products into an electron‐dense core. Retention of cargo by CPE in the immature granule during constitutive‐like secretion also facilitates the trafficking process. CPE can be cleaved in the granule to generate a soluble C‐terminally truncated form of the protein, which has significantly higher specific carboxypeptidase E enzymatic activity than its membrane form. As a soluble protein here, it functions in its classical mode as a carboxypeptidase. Upon secretion of the mature granule, the peptide hormones are released from the core in addition to the soluble CPE and some C terminus‐containing CPE that are extricated from the membrane upon the rapid change in local environment (i.e., pH, Ca2þ, and/or potential raft lipid exchange on the plasma membrane (PM)). The transmembrane form of CPE that remains on the PM is recycled back to the Golgi apparatus by an ARF6‐ dependent endocytic pathway (Arnaoutova et al., 2003b)

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. Figure 3-4 Sequence comparison of PC1, PC2, and CPE. Alignment of the C‐terminal sequences of PC1, PC2, and CPE. Identical residues and chemically similar residues are indicated in boxes

enzyme in cells (Arnaoutova et al., 2003a; Lou et al., unpublished data). The transmembrane domain contains an amphipathic a‐helix associated with lipid rafts at the TGN and this membrane interaction appears to be critical in sorting of PC1/3 to the regulated pathway (Bernard et al., 2003; Lacombe et al., 2005). Recently, Stettler et al. (2005) questioned whether a transmembrane orientation of PC1/3 was possible based on the unlikely nature of the proposed unique transmembrane domain. These authors demonstrated that PC1/3 was not synthesized as a transmembrane protein in COS‐1 cells. Unfortunately, the behavior of transfected PC1/3 constructs in a nonendocrine cell line is not sufficient to conclude categorically a similar behavior for endogenous PC1/3 in an endocrine tissue. PC2 has also been shown to have a transmembrane form and the region consisting of an amphipathic a‐helix (within residues 613–637) has been predicted to be the domain responsible for transmembrane association with lipid rafts (Assadi et al., 2004). Deletion of the last 25 residues at the C terminus of PC2, which includes this predicted transmembrane domain, led to missorting of PC2 in Neuro2a cells, a neuroendocrine cell line, indicating that membrane association with lipid rafts is important for the sorting of PC2. In another study, the PC2 prodomain was shown to bind membranes in vitro. Fumonesin treatment, which disrupts sphingolipid biosynthesis, and site‐directed mutagenesis of the prodomain caused missorting of PC2 in AtT‐20 cells, leading these investigators to conclude that membrane association of the prodomain was important for sorting (Blazquez et al., 2000). It is possible that the prodomain is important in the folding of the enzyme, and disruption in the conformation by site‐directed mutagenesis could prevent lipid‐raft association with the TGN membrane, resulting in constitutive secretion of PC2. As mentioned above, CPE also has an amphipathic a‐helix consisting of residues 411–434, and predicted to be the transmembrane domain (> Figure 3-4). A peptide containing this sequence was shown to insert into artificial model membranes, providing evidence that this domain is responsible for membrane association of CPE (Dhanvantari et al., 2002). Deletion mutation studies indicate that removal of the last 15 amino acids of CPE, which contains a part of the transmembrane domain, caused missorting of CPE to the constitutive pathway in Neuro2a cells (Zhang et al., 2003). Furthermore, treatment of cells with lovastatin, which disrupts cholesterol biosynthesis and hence lipid rafts, prevented sorting of CPE to the secretory granules (Dhanvantari and Loh, 2000). Thus transmembrane association of CPE with lipid rafts at the TGN is critical for sorting to the RSP. Soluble forms of the enzyme can aggregate with the membrane form and be sorted into the RSP. A study on PC5 has shown that there are two forms of the enzyme, one localized in the Golgi apparatus and another in DCGs. Comparison of the two forms revealed that the Golgi form has a truncated C terminus (De Bie et al., 1996). This suggested that sorting information for targeting to the DCGs resides in the C terminus, consistent with the sorting of the other PCs and CPE. In summary, sorting of the proneuropeptide processing enzymes involves interaction with lipid‐ raft microdomains at the TGN. We have proposed that these microdomains are where budding of the TGN membranes occurs to form the secretory granules, since the lipid composition of the granule membranes is highly enriched with cholesterol (70% of total lipid) (Dhanvantari and Loh, 2000). By binding to lipid rafts, PC1/3, PC2, and CPE can be transported to the ISGs as part of the budding membrane.


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Secretory Granule Biogenesis in Neuroendocrine Cells

DCG formation is crucial for the proper sorting, processing, and secretion of neuropeptides and neurotransmitters, as discussed above. DCGs are formed initially as ISGs at the TGN. After processing and condensation of the cargo, they become mature granules and contain an electron‐dense core (Tooze and Stinchcombe, 1992; Tooze et al., 2001). The following sections review the mechanisms involved in DCG biogenesis in neuroendocrine cells.

4.1 Role of Cholesterol in Secretory Granule Biogenesis Cholesterol is a lipid that has been implicated both in the sorting of cargo destined for regulated secretion and in the formation of DCGs (Keller and Simons, 1998; Loh et al., 2004; Tooze, 1998). Cholesterol together with sphingolipids is enriched in detergent‐insoluble, liquid‐ordered membrane microdomains found in the Golgi and the plasma membrane. In 1988, Simons and van Meer proposed that the apical pathway of vesicular trafficking from the TGN (the RSP) was dependent on the presence of cellular cholesterol and on the formation of lipid‐raft microdomains in epithelial cells. In neuroendocrine cells, microdomains are important for creating a rigid environment where proteins can be concentrated via protein–lipid interactions, immobilized, and prepared for packaging into DCGs (Loh et al., 2004). The concept that cholesterol is required for the formation of vesicles from the TGN is just emerging. Although the data are sparse, this section presents evidence supporting a critical role for cholesterol in DCG formation in neuroendocrine cells. Vesicle biogenesis in neuroendocrine cells was recently analyzed in terms of its specific dependence on the presence of cholesterol. By thin‐layer chromotography analysis, cholesterol was revealed to comprise 70% of total granule membrane lipids, which emphasized the importance of cholesterol and cholesterol‐ rich lipids rafts (Dhanvantari and Loh, 2000). It is our hypothesis that this high cholesterol content is derived from the lipid microdomains in the TGN that define the area for budding. Depletion of cellular cholesterol in an endocrine cell line, AtT‐20, resulted in DCGs that could not bud from the TGN (Wang et al., 2000). Upon addition of cholesterol, however, the DCGs formed and were rerouted to the cell periphery (Wang et al., 2000). As mentioned in Sect. 10.3, CPE is a lipid‐raft‐associated processing enzyme in DCGs of neuroendocrine cells. In a study designed to test the possibility that targeting of CPE to the RSP is dependent on the presence of cholesterol, cells were depleted of cholesterol (Dhanvantari and Loh, 2000). CPE was not associated with the detergent‐soluble fraction of membranes; nor was there targeting of this processing enzyme to distal processes, and cells did not respond to regulated secretion. Further, POMC, which binds to CPE for sorting, was also mistargeted, and behaved identically to CPE in secretion studies. These investigations critically introduced cholesterol as a necessary component for budding of granules from the TGN in endocrine cells, and this lipid alone proved to be sufficient to rescue their formation. Mouse models of cholesterol deficiency, the Smith–Lemli–Opitz syndrome (SLOS) and the lathosterolosis mouse, are available (Wassif et al., 2001) and have been analyzed for the role of cholesterol in DCG biogenesis in vivo. The human diseases are caused by a functional lack of 7‐dehydrocholesterol reductase and Sc5d, (Pathosterol desaturase) respectively, the enzymes necessary for the final steps in cholesterol biosynthesis, and patients suffer from multiple congenital malformations (Porter, 2000). In exocrine pancreas, which utilizes the same mechanisms of sorting, budding, and trafficking as neurons and neuroendocrine cells, the SLOS mice displayed severely reduced DCG numbers, highly abnormal granule morphologies, and defective regulated secretion (Gondre´‐Lewis et al., 2006). Consistently, in cholesterol‐ deficient lathosterolosis mice, neuroendocrine islet cells also display a significant decrease in DCG formation (> Figure 3-5). How does cholesterol affect DCG formation at the molecular level? One possible explanation for the phenotype of cholesterol‐deficient mice could be the specificity of cholesterol’s structure. It has been postulated that the structure of cholesterol specifically contributes to the curvature of certain membranes,

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. Figure 3-5 Dense‐core secretory granules (DCGs) in endocrine islet cells. DCG formation is impaired in endocrine islet cells in vivo in the lathosterolosis mouse model of cholesterol deficiency. (a) Control mice have endocrine cells with dense‐core granules of varying size and appearance distributed throughout their cytoplasmic space (bar ¼ 1.0 mm). (b) Mice lacking Sc5d, the enzyme necessary for eventual conversion of lathosterol to cholesterol, show a marked lack of granule formation in pancreatic islet cells (bar ¼ 0.63 mm)

and thus to the budding of vesicles from the TGN. In fact, recent studies have specifically compared cholesterol with other sterols only to reveal that the molecular structure of cholesterol is essential for incorporation in the plasma membrane and promotes the pinching of vesicular structures from the TGN, especially in large vesicles (Bacia et al., 2005; Churchward et al., 2005). For example, substitution of cholesterol with lathosterol or 7‐dehydrocholesterol resulted in increased elasticity and decreased curvature in artificial inverse hexagonal membranes, respectively. Recent work by Bacia and coworkers used giant unilamellar vesicles (GUVs) to demonstrate the dependence on cholesterol for microdomain formation (Bacia et al., 2005). Further, the formation of lipid rafts, the direction and size of curvature, and the ability to form and bud GUVs was critically dependent on the sterol structure and purity of sterol components, consistent with in vivo reports. These studies emphasize the importance of the structural contribution of cholesterol at the initial budding from the TGN in DCG biogenesis.

4.2 Granulogenic Proteins: Assembly Factors It has long been proposed that the formation of DCGs is due to the aggregative property of DCG cargo proteins (Tooze and Stinchcombe, 1992). Many cargo proteins such as granins (e.g., CgA, chromogranin B (CgB), and secretogranin II (SecII)) and prohormones are aggregating under the condition of low pH and high calcium in vitro, similar to the luminal condition of DCGs (Yoo and Albanesi, 1990; Chanat and Huttner, 1991; Colomer et al., 1996; Yoo, 1996; Yoo and Lewis, 1996). In addition, POMC can self‐associate to form oligomers (Cawley et al., 2000). Therefore, it would be plausible that the aggregative nature of cargo would provide a mechanical driving force to form DCGs from the TGN. In support of this idea, overexpression of these DCG cargo proteins (CgA, CgB, SecII, provasopressin, prooxytocin, and POMC) in nonendocrine fibroblasts resulted in the formation of DCG‐like vesicles (Kim et al., 2001; Huh et al., 2003; Beuret et al., 2004). Thus, DCG proteins with aggregative property may function as ‘‘granulogenic’’ proteins to induce the formation of DCGs.


3

4.3 Proteins Regulating DCG Biogenesis Recently, new information has emerged indicating that in addition to the physical formation of DCG through the aggregative properties of the cargo proteins, the number of DCGs formed is highly regulated through a mechanism involving the control of secretory granule protein stability posttranscriptionally and posttranslationally. One of the molecules involved in the regulatory mechanism for DCG biogenesis is CgA. CgA is an acidic glycoprotein highly abundant in DCGs from endocrine, exocrine, and neuronal cells and is considered as a universal marker for these cells (O’Connor et al., 1983; Taupenot et al., 2003). Several physiological functions, including its granulogenic role, as discussed above, have been suggested for CgA and its processed peptides (Taupenot et al., 2003). When neuroendocrine PC12 cells were depleted of CgA expression by antisense RNAs against CgA, DCG formation and regulated secretion was severely impaired in these catecholamine‐secreting cells (> Figure 3-6) (Kim et al., 2001). Reduction of DCG biogenesis was closely correlated with the amount of CgA present in the cells. Similar results were obtained by Yoo and coworkers (Huh et al., 2003), showing significant reduction of DCG formation in PC12 cells treated with siRNA against CgA. Interestingly, CgA deficiency in PC12 cells led not only to the depletion of DCG formation, but also to the degradation of other granule proteins such as CgB, CPE, and synaptotagmin (Kim et al., 2001). Degradation of these proteins was completely recovered by reexpression of CgA (Kim et al., 2001). This observation suggested that CgA was involved in the regulation of DCG formation possibly by controlling stability of granule proteins at the posttranslational level (Kim et al., 2003). Such a role of CgA in DCG formation was also shown in an endocrine cell line, 6T3 cells. These cells, derived from the pituitary corticotroph AtT‐20, lack expression of CgA, DCG formation, and the RSP (Matsuuchi and Kelly, 1991). This missing phenotype was recovered by treatment with 8‐bromo‐cAMP (Day et al., 1995), showing that 6T3 cells possess a potential to revert to apparently normal endocrine cells. In fact, exogenous expression of CgA was able to recover the regulated secretion in these cells (Kim et al., 2001), indicating that CgA was sufficient to reinstate the RSP in 6T3 cells. Consistent with the degradation of granule proteins occurring in CgA‐deficient PC12 cells, exogenously expressed granule protein, CgB was also degraded in 6T3 cells (Kim et al., 2001).

. Figure 3-6 Dense‐core secretory granule (DCG) formation in PC12 cells. Electron micrographs represent wild‐type (WT) PC12 cells and a stable PC12 clone no. 5 expressing antisense CgA RNAs (CgAAS‐5). DCG formation was significantly impaired in CgAAS‐5 cells compared with WT PC12 cells. Arrows indicate DCGs in wild‐type PC12 cells

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Currently, the exact mechanism by which CgA regulates DCG biogenesis in neuroendocrine and endocrine cells is unknown. However, the degradation of granule proteins in cells lacking CgA expression suggests that CgA may regulate DCG formation posttranslationally by preventing degradation of granule proteins in these cells. Supporting this idea, CgA was shown to be a protease inhibitor against IRCM‐serine protease 1 or trypsin (Seidah et al., 1987). It is also possible that CgA or its processed peptides can initiate a signaling pathway to induce expression of protease inhibitors in order to prevent degradation of granule proteins in these cells. Indeed, a processed peptide of CgA, pancreastatin, was shown to initiate a signaling pathway (Gonzalez‐Yanes and Sanchez‐Margalet, 2002). Therefore, CgA by itself or CgA‐mediated activation of gene expression encoding protease inhibitors can potentially regulate the amount of granule cargo proteins available for granule biogenesis and potentially the number of DCGs formed. Indeed, expression of a protease inhibitor, protein nexin-1 (PN-1) has recently been shown to be up-regulated by transfection of CgA into 6T3 cells, which then resulted in an increase in DCG biogenesis (Kim and Loh, 2006). A regulatory mechanism for DCG biogenesis at the posttranscriptional level in pancreatic b‐cells has also been reported. Polypyrimidine‐tract binding protein (PTB) was shown to bind mRNAs encoding granule proteins so as to increase stability of these mRNAs. Upon glucose stimulation in b‐cells, insulin secretion was accompanied by translocation of nuclear resident PTB into the cytoplasm. PTB then bound at the 30 ‐UTR consensus sequence of mRNAs encoding a receptor‐tyrosine‐phosphatase‐like protein (ICA512/ IA2), insulin, PC1/3, PC2, and CgA to stabilize these transcripts, resulting in increased translation of these proteins in the b‐cells. Downregulation of PTB by siRNA resulted in the loss of DCG formation as well as the decrease of insulin, CgA, SecII, PC1/3, PC2, synaptobrevin 2, and synaptophysin (Knoch et al., 2004). These observations supported a role of PTB in a posttranscriptional mechanism to increase the level of DCG proteins and DCG biogenesis in pancreatic b‐cells upon stimulation by glucose. Regulation of DCG biogenesis appears to be mediated by proteins (PTB and CgA) involved in stabilization of DCG mRNAs and proteins at the posttranscriptional and posttranslational levels. These regulators ultimately determine the levels of DCG proteins and the quantity of DCGs formed in neuroendocrine and endocrine cells.

5

Conclusion

Current evidence indicates that proneuropeptides and BDNF are sorted actively to the regulated pathway by interaction with a sorting receptor, such as CPE, while the sorting of their processing enzymes, PCs and CPE, appear to be mediated by interaction of their C terminus with lipid rafts at the TGN. As the field progresses, other sorting receptors or membrane‐binding proteins that are involved in cargo protein sorting into the RSP are likely to emerge. Furthermore, besides cholesterol and CgA, we anticipate that additional lipid and protein components, which are essential for DCG formation, as well as the mechanisms involved in regulating quantitative DCG biogenesis in different types of secretory cells will also be identified in the future.

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Protein Synthesis at Synaptic Sites on Dendrites

O. Steward

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170

2

Form Implies Function: Morphology of Protein Synthetic Machinery at Synapses . . . . . . . . . . . 171

3

The Spine as a Protein Synthetic Compartment: Spatial Constraints Due to Size . . . . . . . . . . . 172

4

Documentation of Local Protein Synthesis in Dendrites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173

5

Presence of Elements of the ER and Golgi Apparatus in Dendrites . . . . . . . . . . . . . . . . . . . . . . . . . . . 174

6 6.1 6.2 6.3 6.4

What mRNAs Are Present in Dendrites/What Proteins Are Synthesized Locally at Synapses? . Assessment of Subcellular Fractions Containing Pinched‐Off Dendrites . . . . . . . . . . . . . . . . . . . . . . . In Situ Hybridization Analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Isolation and Amplification of mRNA from Neurites of Neurons Growing in Culture . . . . . . . Microarray Studies of mRNA from Dissected Dendritic Laminae In Vivo . . . . . . . . . . . . . . . . . . . . .

7

Local Synthesis of Components of Multimolecular Signaling Complexes: Is There a Cotranslational Assembly? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180

8 8.1 8.2 8.3 8.4

Dendritic Transport of RNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Messenger RNA Transport Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Cis‐Acting Sequences That Identify mRNA for Dendritic Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Dendrite‐Specific RNA Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182 Mechanisms Underlying mRNA Localization at Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183

9 9.1 9.2 9.3 9.4 9.5 9.6

Regulation of Translation of Dendritic mRNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 Translational Repression of Localized mRNAs in Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184 Translational Repression of Dendritic mRNAs While They Are in Transit . . . . . . . . . . . . . . . . . . . . . 184 The Fragile‐X/BC1 Connection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 Micro‐RNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186 Regulation of Translation by Synaptic Signals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186 Regulation of Protein Synthesis in Developing Dendrites by Miniature Synaptic Events . . . . . . 187

174 175 175 179 180

10 Protein Synthesis in Axons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 10.1 Protein Synthesis in Growing Axons and Growth Cones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 188 11

#

Degradation of Dendritic mRNAs and Locally Synthesized Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . 189


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4

Protein synthesis at synaptic sites on dendrites

Abstract: An important aspect of gene expression in neurons involves the delivery of mRNAs to different intracellular destinations where certain proteins can be locally synthesized. In mature neurons, local protein synthesis at synaptic sites on dendrites is now known to play a key role in activity‐dependent synaptic modifications including long‐term potentiation (LTP) and long‐term depression (LTD). In developing neurons, local synthesis in the growth cone is important for extension and guidance. Here, we summarize some of the key aspects of local protein synthesis in dendrites, focusing especially on how local translation of mRNAs is regulated by synaptic activity. List of Abbreviations: BDNF, brain-derived neurotrophic factor; CAMKII, calcium-calmodulin dependent protein kinase II; CPEB, cytoplasmic polyadenylation element-binding protein; CPE, cytoplasmic polyadenylation element; ER, endoplasmic reticulum; FMRP, fragile X mental retardation protein; GFP, green fluorescent protein; GluR, glutamate receptor; IEG, immediate early gene; INSp3, inositol trisphoaphate; IRES, internal ribosome entry site; LTD, long term depression; LTP, long term potentiation; MAP2, microtubule-associated protein 2; MBP, myelin basic protein; mRNA, messenger RNA; mTOR, mammalian target of rapamycin; NMDAR1, N-methyl-D-aspartate receptor I; NRC, NMDA receptor complex; RER, rough endophasmic reticulum; RNA, ribonucleic acid; rRNA, ribosomal RNA; RTPCR, real time polymerase chain reaction; SPRC, synapse-associated polyribosome complexes; SRP, signal recognition particle; TTX, tetrodotoxin; UTR, untranslated region; VSVG, vesicular stomatitis virus glycoprotein

1

Introduction

A central tenet of neuronal cell biology is that most of the protein synthetic machinery in neurons is localized in the perinuclear cytoplasm in the neuronal cell body, and that most of the proteins required for axons and dendrites are transported to their final destinations. Beginning in the early 1980s, however, evidence began to emerge that there were also outposts of protein synthetic machinery at different intracellular locations, especially synaptic sites on dendrites, as well as in developing axons and growth cones. The story line on dendritic protein synthesis was launched by the discovery that polyribosomes and associated membranous cisterns were selectively positioned beneath and within dendritic spines (Steward and Levy, 1982; Steward, 1983; Steward and Fass, 1983). We now term these synapse‐associated polyribosome complexes (SPRCs). The selectivity of the localization suggested several hypotheses, including: (1) that SPRCs synthesized key molecular constituents of the synapse, (2) that local translation was regulated by signaling events at the synapse, and (3) that local translation played a key role in synapse plasticity. These hypotheses formulated in 1982 have been confirmed and extended by a host of studies (Steward and Schuman, 2001, 2003). Running in parallel with the story of local protein synthesis in dendrites was a story line that developed in fits and starts regarding local protein synthesis in axons. There has been general agreement that protein synthesis occurs in invertebrate neurites (Alvarez et al., 2002), but whether similar mechanisms existed in vertebrate axons remained controversial. Recent studies, however, have established the existence of protein synthetic machinery and mRNAs in growing vertebrate axons, especially growth cones, and demonstrated that this machinery and the local protein synthesis it allows plays a key role in growth cone function (Steward, 2002a). Even more recent studies have provided evidence that there is also local protein synthesis in at least some mature axons, and that this local synthesis plays a critical role in signaling from the axon to the nucleus in situations involving axonal injury (Zheng et al., 2001; Hanz et al., 2003). Here, we summarize what is known about the mechanisms for local protein synthesis at synaptic sites on dendrites. We focus here on cell biological issues, and mention only briefly the role of local synthesis in synaptic plasticity, which has been considered in other recent reviews (Steward and Schuman, 2001, 2003; Schuman et al., 2006).


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4

Form Implies Function: Morphology of Protein Synthetic Machinery at Synapses

An important clue about the function of SPRCs is their highly selective positioning beneath synapses. At spine synapses, SPRCs are most often localized at the base of the spine in the small mound‐like structures from which the neck of the spine emerges, and less frequently in the spine neck and head (Steward and Levy, 1982). Interestingly, following synaptic stimulation leading to long‐term potentiation (LTP) in the CA1 region of the hippocampus, polyribosomes are more numerous in the spine proper, suggesting that they can shift position as a result of signals generated at the synapse (Ostroff et al., 2002). At nonspine synapses (both excitatory and inhibitory) SPRCs are clustered beneath the postsynaptic membrane specialization (Steward et al., 1996). SPRCs, located beneath the synapse, are ideally situated to be influenced by ionic and/or chemical signals from the synapse as well as by events within the dendrite proper. An important implication of this selective localization is that there must be some mechanism that causes ribosomes, mRNA, and other components of the translational machinery to localize selectively in the postsynaptic cytoplasm. The details of the mechanisms underlying this selective localization remain to be established. The proportion of synapses with underlying SPRCs varies by cell type, position of the synapse on the dendrite, and developmental age. Reconstructions of dendrites in the dentate gyrus of adult rats reveal that about 25% of the spine synapses on mid proximo‐distal dendrites have underlying polyribosomes (Steward and Levy, 1982). The incidence is higher at synapses on proximal dendrites, where many spines have multiple polyribosome clusters. Studies of ribosomes (not polyribosomes) in serially reconstructed spines revealed that most spines on pyramidal neurons in the cerebral cortex contained ribosomes, whereas the incidence of ribosomes was lower in spines on cerebellar Purkinje cells (Spacek and Hartmann, 1983). Polyribosomes are especially prominent during periods of maximal synaptogenesis, when most synapses have underlying polyribosomes, and many synapses have multiple clusters. These facts imply that local synthesis is especially important during periods of synapse growth (Steward and Falk, 1986). The fact that polyribosomes are present at some, but not all, synapses in mature animals raises the question of whether SPRCs can shuttle from one synapse to another, or whether the presence of the machinery marks synapses that are capable of, and in the process of local protein synthesis. The fact that polyribosomes appear to move from the base of the spine into the spine head (Ostroff et al., 2002) indicates that polyribosomes can shift position within an individual synapse, but leave open whether polyribosomes can move from one synapse to another. Also unknown is whether a given polyribosome moves as a unit, or whether the individual ribosomes dissociate from the polyribosome, move, and then reinitiate on some other mRNA. One important feature of SPRCs is that the number of ribosomes at individual synaptic sites is quite limited. If ultrastructural studies are actually identifying all dendritic ribosomes (including unassociated ribosomal subunits), then the number of ribosomes in a polyribosome at a typical synapse in the hippocampus and dentate gyrus ranges from about 3–28 with the average being 8 (Steward and Reeves, 1988; Ostroff et al., 2002). Most synapses have only one or two polyribosomes, and some synapses apparently have none. Unquestionably, the availability of ribosomes must constrain the overall capacity for protein synthesis at individual synapses. It is not known how this limited machinery is allocated to the different mRNAs that are potentially available for translation. Also unknown is whether the translational machinery is running at full capacity. When ribosomes are present in clusters (polyribosomes) it is presumed that they are translating a single mRNA molecule. If this assumption holds at synapses, and given that there are usually no more than one or two clusters present at a typical spine synapse, it follows that one or two mRNAs are being translated at a given time. Indeed, looking at reconstructions of synapse‐associated polyribosomes or even individual electron micrographs, one can easily recognize the ‘‘string of pearl’’ configurations, suggesting assembly onto a single mRNA molecule (Ostroff et al., 2002). If all ribosomes are so engaged, then the overall rate of

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translation can only be enhanced by increasing the scanning rate or the rate of addition of amino acids to the nascent peptide. If all ribosomes are already engaged, it follows that changing the initiation probability (for example as a result of phosphorylation of initiation factors) would probably not increase overall translational capacity. Conversely, decreasing translation initiation could decrease protein synthetic rate, and a resulting decrease in the number of polyribosome clusters would be expected. An alternative, which is unlikely based on cell biological orthodoxy, is that each individual ribosome is engaged with a different mRNA. In this case, there could be a greater number of different proteins being synthesized at the same time, but the total number of individual ribosomes would still be limiting for how many mRNAs could be translated at a given time. Moreover, as discussed further below, considerations of size constraints suggests that it would be difficult to pack very many mRNA molecules into the small area beneath a synapse. So, as in the model above, it is hard to imagine how there could be meaningful increases in translational capacity as a result of synaptic signals if all the ribosomes are already engaged. A third alternative is that a proportion of the ribosomes at a synaptic site are unengaged in the absence of some stimulus (for example a stimulus that triggers plasticity), and that signals from the synapse trigger initiation. This is the only mechanism that seems likely to result in a meaningful increase in overall translational capacity at an individual synapse without increasing the number of available ribosomes. A way around the numbers problem would be if additional ribosomes could be rapidly recruited to individual synaptic sites. As noted above, there is some evidence for shifts in ribosome position within spines following stimuli that induce LTP (Ostroff et al., 2002). To increase overall translational capacity at one synapse, however, the ribosomes would have to come from somewhere else, which would presumably decrease translational capacity at nearby synapses. Thus, again, the overall translational capacity of small dendritic segments would presumably be rather stable unless there were significant numbers of unengaged ribosomes or a capacity to transport additional ribosomes from a distance. Finally, it is important to mention the possibility of local biogenesis of ribosomes. The cell biological orthodoxy is that ribosomes are assembled in the nucleolus from rRNA and ribosomal proteins, but recent studies have demonstrated that mRNAs for certain ribosomal proteins are present in neurites of invertebrate neurons (Moccia et al., 2003). The proteins encoded by mRNAs in neurites tend to be located physically at or near the edge of the ribosome where the two ribosomal subunits link up, and so local synthesis of these subunits could promote assembly of the subunits into a ribosome. Nevertheless, this model still requires that nearly complete ribosomal subunits be present to which the newly synthesized proteins can be added. It is not clear whether it is possible to see ribosomal subunits at the electron microscopic level. If ribosomal subunits are visible using routine electron microscopy, then there clearly is not an excess of subunits surrounding synapses, which could be recruited into polyribosomes. If subunits are invisible by standard methods, then assumptions based on ribosome numbers will have to be modified. It is also not known whether mRNAs for ribosomal proteins are present in dendrites of mammalian neurons in vivo.

3

The Spine as a Protein Synthetic Compartment: Spatial Constraints Due to Size

One additional point to consider is the size of the mRNAs present in dendrites. If mRNA was in a configuration similar to the B‐form of DNA, then the overall length of an mRNA can be estimated as 0.34 nm per base multiplied by the number of bases. Considering representative mRNAs that are prominent in dendrites (see below), the mRNAs for CAMKII, Arc, BDNF and MAP2 would be about 1.57, 1.03, 1.32, and 3.17 mm long, respectively (the latter is longer than the average spine). > Figure 4-1 illustrates how these sizes compare to the size of typical spine synapses, other dendritic organelles, and the estimated size of putative RNA granules. Of course the mRNAs would not be straightened out, as shown, but would presumably be configured in tangled loops and hairpins. Still, even when in their tangled three‐dimensional form, mRNAs must occupy considerable space, especially if they are associated with mRNA‐binding proteins. These sizes illustrate that the area beneath and within spines may be very crowded if more than one mRNA is being translated at any given time. Adding a spine apparatus in, which is present at many


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. Figure 4-1 Approximate sizes of representative dendritic mRNAs and translational elements at synaptic sites on dendrites. The approximate size range of spine synapses that would be found in rat forebrain structures like the hippocampus and cerebral cortex is depicted. The lines represent the approximate length of representative dendritic mRNAs if they were straightened out. Shading indicates the length and position of the coding region (from Schuman et al., 2006)

synapses, further limits the space that would be available for shifting mRNAs around in the very limited cytoplasmic microcompartment beneath an individual synapse. The extent to which such spatial limitations actually constrain translation remains to be defined.

4

Documentation of Local Protein Synthesis in Dendrites

The presence of polyribosomes implies active protein synthesis, but it was important to demonstrate protein synthetic activity directly. To definitively establish that protein synthesis is indeed occurring in local compartments, be it dendrites or axons, the cell body must be ruled out as a protein synthesis source. This is important because protein synthetic machinery is highly concentrated in neuronal somata and given rates of rapid transport, newly synthesized proteins can be delivered from cell bodies throughout dendrites within minutes. Initial evidence for local synthesis in dendrites came from studies using subcellular fractions of synaptodendrosomes or synaptoneurosomes, but these fractions are usually (probably always) contaminated to some degree by fragments of cell bodies and glia, which cannot be excluded as the source of protein synthetic activity being detected. The development of a novel cell culture platform that allowed physical isolation of dendrites made it possible to demonstrate a capacity for local incorporation of amino acid precursors in dendrites using autoradiographic techniques (Torre and Steward, 1992), but limitations on the resolution of autoradiography and the likelihood that newly synthesized proteins moved made it impossible to define the actual sites of protein synthesis in the isolated dendritic segments. These limitations have been largely overcome in recent studies that use physically and optically isolated dendrites and reporter constructs made up of GFP‐tagged proteins modified in such a way that they are relatively immobile after synthesis. Imaging the GFP constructs in living dendrites has then allowed for definitive analyses of the sites and the nature of dendritic protein synthesis (Aakalu et al., 2001; Job and Eberwine, 2001). These studies have revealed persistent hotspots for protein synthesis along the length of the dendrite that were located near ribosomes and synapses, lending further support to the idea that dendritic sources of protein synthesis may subserve a small synaptic domain. As discussed further below, live‐cell imaging methods have also begun to provide insights into the signal transduction processes that regulate protein synthesis in response to synaptic signals.

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Presence of Elements of the ER and Golgi Apparatus in Dendrites

Integral membrane proteins (receptors for example) and proteins for release are synthesized by the rough endoplasmic reticulum (RER) and are usually glycosylated. Thus, an important issue has been whether ribosomes are present on membranes in an RER‐like configuration, and whether ER and Golgi apparatus enzymes are present that could mediate posttranslational modifications. Reconstructions of dendrites of dentate granule cells and hippocampal pyramidal cells indicate that about 50% of the polyribosomes beneath synapses are associated with tubular cisterns, suggesting that the SPRC/cisternal complex may be a form of RER (Steward and Reeves, 1988). Later immunocytochemical studies revealed the presence of different markers of the RER in dendrites of neurons in culture, including ribophorin I (Torre and Steward, 1996), the signal sequence receptor TRAPP, and the signal recognition particle (SRP) that directs nascent polypeptide chains to the RER (Tiedge and Brosius, 1996). Electron microscopic immunocytochemical studies also revealed that the membranous cisterns present near spine synapses stain for the Sec6Ia protein complex, which is part of the machinery for translocation of proteins through the RER during their synthesis (Pierce et al., 2000). Moreover, dendrites that have been separated from their cell bodies incorporate sugars in a tunicamycin‐sensitive fashion (Torre and Steward, 1996) and are capable of translating membrane protein‐encoding mRNAs (glutamate receptors) and inserting these into the plasma membrane (Kacharmina et al., 2000). Together, these studies indicate that elements of RER and Golgi apparatus are present in dendrites in a configuration that can support synthesis, posttranslational processing, and membrane insertion of locally synthesized proteins. In nonneuronal cells, secretory and membrane proteins are synthesized on the RER, packaged into post‐ER carriers (coated vesicles), and transported to a centrally located Golgi apparatus, where protein processing and sorting occurs. Live‐cell imaging using a GFP‐tagged temperature‐sensitive mutant of the vesicular stomatitis virus glycoprotein (VSVG), and ER and Golgi apparatus markers, has documented the presence of functional ER, post‐ER carriers, Golgi apparatus elements, and post‐Golgi apparatus trafficking in dendrites (Horton and Ehlers, 2003). VSVG–GFP moves from ER elements in dendrites into highly mobile tubero‐vesicular structures, and the tubero‐vesicular structures then move bidirectionally in dendrites, fusing with stationary structures that stained for Golgi apparatus markers (GM130). The accumulation of VSVG–GFP into tubero‐vesicular structures occurred at defined immobile foci that were positive for Sec13, a marker of ER exit sites in other cells. In the context of local synthesis, it is noteworthy that VSVG–GFP‐containing tubero‐vesicular structures were transported bidirectionally for tens of micrometers. This raises a puzzling issue. Why go to the trouble of distributing protein synthetic machinery throughout dendrites if the proteins produced are rapidly transported to other locations? Perhaps VSVG–GFP is missing some critical address tag that is possessed by proteins locally synthesized that would target the proteins to the nearby synapse. Alternatively, perhaps our ideas about the purpose of local synthesis still need fine‐tuning. There is still uncertainty about the organelle responsible for Golgi apparatus‐like activity in dendrites. Electron microscopic immunocytochemical studies indicate that certain Golgi apparatus marker proteins are present in cisterns in spines (Pierce et al., 2000). On the other hand, the highly mobile Golgi apparatus elements identified by live‐cell imaging are localized in the dendritic shaft and are not associated with synapses (Horton and Ehlers, 2003). In either case, the spine apparatus, which has a form that has invited speculation that it might be a mini‐Golgi apparatus (Steward and Reeves, 1988), has not been implicated in Golgi apparatus function. Thus, some 50 years after its discovery, the function of the spine apparatus and the related cisternal apparatus at nonspine synapses remains a mystery.

6

What mRNAs Are Present in Dendrites/What Proteins Are Synthesized Locally at Synapses?

Three main approaches have been used to try to identify mRNAs present in dendrites and the proteins that are locally‐synthesized: (1) assessments of subcellular fractions that contain pinched off dendrites, (2) in situ hybridization analyses in brain tissue and neurons growing in culture, and (3) isolation and


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amplification of mRNA from neurites of neurons growing in culture. The type of information provided by each approach differs, and each has its associated caveats and limitations.

6.1

Assessment of Subcellular Fractions Containing Pinched‐Off Dendrites

Subcellular fractions called ‘‘synaptodendrosomes’’ (Rao and Steward, 1991) or ‘‘synaptoneurosomes’’ (Weiler, 1991; Weiler and Greenough, 1993; Weiler et al., 1997), depending on the techniques used, have been used for biochemical studies of proteins locally synthesized (Rao and Steward, 1991; Leski and Steward, 1996; Zalfa et al., 2003), and to isolate and identify mRNAs (Rao and Steward, 1991; Chicurel et al., 1993; Bagni et al., 2000). A major caveat of this approach, however, is that the fractions may be contaminated with fragments of neuronal and glial cell bodies. For example, some such fractions contain high levels of the mRNA encoding glial fibrillary acidic protein, indicating contamination by glial cells (Rao and Steward, 1991; Chicurel et al., 1993). There have been continuing efforts to refine subcellular fractionation approaches so as to yield more pure fractions. In this regard, one recent study reported a fractionation approach that yields synaptosomes in which GFAP mRNA is not detected by RT/PCR (Bagni et al., 2000), suggesting a lack of contamination by glial fragments. Previously identified dendritic mRNAs for CAMKII, Arc, and an inositol‐1,4,5‐trisphosphate receptor (InsP3R1) were detected in the fractions, and also the mRNA for fragile X mental retardation protein (FMRP) (see below). These fractions may represent a more pure population of synaptodendrosomes than have been available previously, which could provide a means to identify novel dendritic mRNAs, but it is important to assess each individual preparation for purity. Even with this control, concerns about contamination still cannot be completely dismissed.

6.2

In Situ Hybridization Analyses

Probably the most conservative approach to identifying mRNAs in dendrites has been to use in situ hybridization (by conservative, we mean the least potential for false‐positive findings). In tissue sections, dendritic localization of mRNAs is inferred by the pattern of labeling in brain regions where there are distinct neuropil layers, which contain dendrites but few neuronal cell bodies (cortical regions including the hippocampus and the cerebellar cortex). It is important to establish that the mRNA is in fact present in dendrites and not in glial cells, which can be done using nonisotopic in situ hybridization techniques. > Table 4-1 lists mRNAs that show substantial dendritic localization in vivo (i.e., labeling that extends for several hundred mm from the cell body). The list includes mRNAs encoding cytoplasmic, cytoskeletal, integral membrane, and membrane‐associated proteins that have quite different functions. The different mRNAs are expressed differentially by different neuron types, and exhibit somewhat different localization patterns within dendrites. Some of the mRNAs (the mRNA for calmodulin for example) are present in dendrites during early development, and absent in mature neurons (Berry and Brown, 1996). The table does not include mRNAs that extend for only a few tens of micrometers into proximal dendrites (for example mRNAs for the protein kinase C substrates GAP43 and RC3) (Laudry et al., 1993). Data from in situ hybridization analyses have revealed several important conclusions. First, it is now clear that there are actually multiple patterns of mRNA distribution within dendrites. For example, certain mRNAs are present throughout the full extent of the dendritic arbor including the mRNAs for CAMKII, dendrin, the IP3 receptor, neuralized, and Arc when it is induced by a seizure. Other mRNAs are concentrated in particular dendritic segments, including the mRNA for MAP2, which is concentrated in the proximal 1/3–1/2 of the dendrite and is absent from distal dendrites. The mRNA for EF1a is also concentrated in laminae associated with particular afferent systems, although it is also present throughout dendrites at lower levels (Huang et al., 2005). The fact that different mRNAs are localized in different dendritic domains means there must be several different localization signals in the mRNA and different docking mechanisms (the term ‘‘docking’’ refers generically to whatever exists in particular dendritic domains that causes particular mRNAs to accumulate there). Second, different neuron types have a different complement of

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Cytoplasmic RNA binding

Cytoskeleton‐associated activity‐ induced NRC/psd protein

Mainly proximal 1/3

Throughout (when induced)

Forebrain

Widespread

Widespread

Widespread but at highest levels in interneurons

Cortex, hippocampus, dentate gyrus depending on inducing stimulus

EF1an

Shank AKA SSTRIPo

FMR1/fragile X mental retardation protein FMRPp Activity‐regulated genes Arc/Arg 3.1q

Throughout

Throughout

NRC/psd protein

Cytoplasmic

NRC/psd protein

Cytoplasm and membrane‐ associated Integral membrane Integral membrane Soluble Cytoskeletal Cytoplasmic Chromodomain‐containing Membrane associated

Membrane‐associated

Membrane‐associated NRC/psd protein Putative membrane

SAP90/SAPAPm

Proximal‐middle? Proximal Proximal‐middle Proximal‐middle Throughout Proximal 1/3–1/2 Throughout but most abundant in proximal 1/3 Throughout

Proximal‐middle

Throughout

Throughout

Throughout

Class of protein Cytoskeletal

NMDAR1f Glycine receptor a‐subunitg Vasopressinh Neurofilament protein 68i Neuralizedj MRG15k EFA6Al

Calmoduline

G protein g subunitd

Dendrinc

CAMII kinase a subunit b

Localization in dendrites Proximal 1/3–1/2

Cell type Cortex, hippocampus, dentate gyrus Cortex, hippocampus, dentate gyrus Hippocampus, dentate gyrus, cerebral cortex Cortex, hippocampus, dentate gyrus, striatum Cortex, hippocampus, Purkinje cells Dentate gyrus Motoneurons Hypothalamo‐hypophyseal Vestibular neurons Widespread Widespread Forebrain

Actin binding??

psd95‐associated protein Elongation factor actin‐interacting Links InsP3 to GKAP /psd95 Translation repression?

Metabotropic receptor signaling Ca2þ signaling in conjunction with CAMII kinase Receptor Receptor Neuropeptide Neurofilament Transcription repressor Nucleolar? ADP‐ribosylation factor

Multifunctional kinase Ca2þ signaling Unknown

Protein function Microtubule‐associated

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mRNA MAP2a

. Table 4-1 mRNAs that have been shown to be localized within dendrites of mature neurons in vivo by in situ hybridization. Not shown are mRNAs that are localized only in the most proximal segments

176 Protein synthesis at synaptic sites on dendrites

Purkinje cells

Purkinje cells

Purkinje cells

Genes expressed primarily in Purkinje cells InsP3 receptors

L7t

PEP19s

b

Garner et al. (1988) Burgin et al. (1990) c Herb et al. (1997) d Watson et al. (1994) e Berry and Brown (1996) f Benson (1997) g Racca et al. (1997) h Prakash et al. (1997) i Paradies and Steward (1997) j Timmusk et al. (2002) k Matsuoka et al. (2002) l Sakagami et al. (2004) m Welch et al. (2004) n Huang et al. (2005) o Zitzer et al. (1999) p Ferrari, F., Huang, F., Bagni, C., and Steward, O in preparation. q Lyford et al. (1995); Link et al. (1995) r Tongiorgi et al. (2005) s Furuichi et al. (1993) t Bian et al. (1996)

a

Hippocampus/limbic system

BDNFr

Proximal 1/3

Throughout

Throughout

Throughout? (when induced by epileptogenic events)

Cytoplasmic

Integral membrane (endoplasmic reticulum) Cytoplasmic?

Soluble

Homology to c‐sis PDGF oncogene Signaling? Ca2þ‐binding

Ca2þ signaling

Growth and trophic factor


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dendritic mRNAs. The mRNAs most abundant in Purkinje neurons (mRNAs for the IP3 receptor and L7) are present at low levels or absent from neurons in the forebrain. Conversely, many of the mRNAs prominent in forebrain neurons (CAMKII and Arc) are not present or are present at very low levels in Purkinje neurons. Importantly, few mRNAs have yet been identified in neurons in the thalamus, brainstem, and spinal cord, although neurons in these regions clearly have SPRCs. Third, certain mRNAs are present in dendrites in developing but not in mature animals suggesting that neurons may use mRNA sorting for different purposes at different times in their life history. Fourth, at least two dendritic mRNAs (Arc and BDNF) are induced by specific patterns of synaptic activity and then transported into dendrites (see below). The mRNA for FMRP requires special mention because initial studies using radioactive in situ hybridization techniques failed to detect FMRP mRNA in dendrites in vivo (Hinds et al., 1993; Valentine et al., 2000), despite the fact that FMRP mRNA can be detected (using RT‐PCR) in isolated synaptodendrosomes (Zalfa et al., 2003). Recently, however, studies using sensitive nonisotopic in situ hybridization techniques have shown that FMRP mRNA is detectable in dendrites of neurons in vivo (Ferrari et al., 2005 and in preparation). In most neurons, levels of dendritic labeling were quite low. Interestingly, however, labeling was prominent in the dendrites of inhibitory interneurons in the hippocampus, and these same interneurons also expressed high levels of the nontranslated pol‐3 transcript BC1, which is also localized in dendrites. This is of particular interest because of recent evidence that BC1 links particular mRNAs to FMRP, and plays a role in repressing translation (Zalfa et al., 2003). This ties in nicely with other work implicating BC1 as a regulator of translation initiation of dendritic mRNAs (Wang et al., 2002). These findings have led to the provocative idea that the loss of FMRP in fragile X mental retardation syndrome could lead to a dysregulation of mRNA translation at synapses (Zalfa et al., 2003). Of special note are mRNAs that are induced by neuronal and/or synaptic activity and then delivered into dendrites. The prototype of activity‐induced dendritic mRNAs is Arc (activity‐regulated cytoskeleton‐ associated protein) (Lyford et al., 1995) also known as Arg 3.1 (Link et al., 1995). Arc was initially discovered in screens for novel immediate early genes (IEGs), defined in this case as genes induced by activity in a protein synthesis‐independent fashion. Like other IEGs, Arc expression is strongly induced by neuronal activity. However, in dramatic contrast to the mRNAs of other IEGs, Arc mRNA is rapidly delivered into dendrites. The fact that Arc mRNA is rapidly delivered into dendrites whereas the mRNAs for other immediate early genes remain tightly localized within the cell body indicates that sorting of newly synthesized mRNAs is a highly regulated process (Wallace et al., 1998). The most striking feature of Arc is that it localizes selectively at activated synaptic sites (Steward et al., 1998b). This was demonstrated in studies in which Arc was induced by stimulating the perforant path in vivo, which terminates in a topographically organized fashion along the dendrites of dentate granule cells. Following stimulation that induces LTP (400 Hz trains, 8 pulses per train, delivered at a rate of 1/10 s), Arc expression is strongly induced, and Arc mRNA is transported into dendrites. If synaptic activation continues as the mRNA is delivered into dendrites, the newly synthesized mRNA accumulates selectively in exactly the location of the band of synapses that had been activated. Activation of other afferent systems that terminate at different locations on the dendrite causes the newly synthesized mRNA to localize selectively in the activated dendritic domains (Steward et al., 1998b). Subsequent studies revealed that both induction of transcription and targeting of Arc mRNA to active synapses depended on NMDA receptor activation (Steward and Worley, 2001b). As discussed in previous reviews, Arc has a number of features that would be consistent with its key role in the enduring forms of synaptic modification that require new gene expression (Steward and Worley, 2001a; Steward, 2002a). Moreover, it has been shown that knocking down Arc expression with antisense nucleotides disrupts hippocampal LTP and memory consolidation (Guzowski et al., 2000). Moreover, recent studies have provided insights into the actual functional role of Arc protein at the synapse (for an overview, see (Tzingounis and Nicoll, 2006)). Specifically, it has been found that Arc protein interacts with endocytic machinery at the synapse to regulate AMPA receptor endocytosis, causing a down-regulation of AMPA receptor numbers in the postsynaptic membrane (Chowdhury et al., 2006; Rial Verde et al., 2006; Shepherd et al., 2006). These results, together with studies of mice lacking Arc (also called Arg 3.1)


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(Path et al., 2006) indicate that Arc protein plays a role in re-scaling synaptic strength during the consolidation phase of memory. Recent studies have revealed that BDNF mRNA has some of the same characteristics as Arc. Like Arc, BDNF mRNA is induced by seizures and intense synaptic activity. There is evidence that BDNF mRNA is transported in dendrites of young neurons in culture (Tongiorgi et al., 1997), but studies in vivo had previously failed to reveal evidence that BDNF mRNA was delivered into mature dendrites even when it was strongly induced by certain types of seizures (Isackson et al., 1991). Recent studies have revealed, however, that BDNF mRNA is delivered into dendrites in the first few hours after ‘‘epileptogenic’’ stimuli that cause the development of a circuit that later exhibits a propensity to seizures (Tongiorgi et al., 2005). BDNF mRNA and protein accumulate in dendrites in all hippocampal subfields after pilocarpine seizures, and in selected subfields after other epileptogenic stimuli (kainate and kindling). Interestingly, BDNF accumulates selectively in discrete dendritic laminae (for example, the inner molecular layer of the dentate gyrus), suggesting targeting to synapses that are active during seizures. Dendritic targeting of BDNF mRNA was not seen after intense stimuli that are nonepileptogenic (electroconvulsive seizures and high‐frequency stimulation). These results suggest that dendritic accumulation of the BDNF mRNA and protein play a critical role in the cellular changes leading to epilepsy (i.e., in the process of epileptogenesis). It remains to be seen whether BDNF is selectively targeted to active synapses, and if so, what signal transduction cascades mediate this targeting. In situ hybridization data indicate that several other mRNAs are present constitutively in the dendrites of young neurons developing in vitro, including mRNAs for BDNF and trkB receptors (Tongiorgi et al., 1997), the mRNA for a fatty‐acylated membrane‐bound protein called ligatin (Severt et al., 2000), and the mRNA for b‐actin (Tiruchinapalli et al., 2003). The mRNAs are usually present in granules, and live‐cell imaging has documented the movement of mRNA‐containing granules within dendrites (Knowles et al., 1996). The extent of dendritic labeling and movement of mRNA‐containing granules into dendrites has been shown to be enhanced by neurotrophin treatment (Knowles and Kosik, 1997) and depolarization by KCl (Tiruchinapalli et al., 2003), indicating that the distribution of mRNA in the dendrites of neurons in culture may be dependent in part on different growth substrates and signaling molecules. Despite prominent dendritic labeling in cultured neurons, the mRNAs for BDNF and trkB receptors appear to be largely restricted to the region of the cell body in young neurons in vivo (Dugich et al., 1992). The same is true of ligatin (Perlin et al., 1993). These results raise the possibility that neurons developing in culture have a different complement of dendritic mRNAs than their counterparts in vivo. As noted above, the mRNA for BDNF is transported into the dendrites of mature neurons in vivo in response to signals that trigger plastic changes involved in epileptogenesis (Tongiorgi et al., 2005), raising the possibility that differences in mRNA distribution between neurons in culture and neurons in vivo may be due to differences in activity or the overall state of the neuron.

6.3

Isolation and Amplification of mRNA from Neurites of Neurons Growing in Culture

Only a small number of mRNAs are detectable in dendrites by in situ hybridization analyses, whereas the vast majority of mRNAs that have been evaluated are localized exclusively in neuronal cell bodies. A different approach, however, has provided evidence for a large and heterogeneous complement of dendritic (or at least neuritic) mRNAs in neurons in culture. Miyashiro and coworkers (1994) used patch pipettes to aspirate the cytoplasmic contents of individual neurites of hippocampal neurons in culture and then used RNA amplification techniques and mRNA expression profiling. A large number of different mRNAs were detected, but many of these cannot be detected in dendrites by in situ hybridization analyses. It is possible that the amplification techniques detect mRNAs that are not accessible to probes in situ. Alternatively, the amplification techniques are extremely sensitive, and may detect mRNAs that are present at levels below the threshold for detection by in situ hybridization. If the latter interpretation holds, the issue arises of what level of mRNA is necessary to generate biologically significant amounts of protein.

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Microarray Studies of mRNA from Dissected Dendritic Laminae In Vivo

One recent study has used microarray analyses to identify mRNAs that were enriched in dendritic laminae in vivo (specifically, the stratum radiatum of the hippocampus). In all, 154 candidate dendritic mRNAs were identified and were annotated and sorted into functional groups (Zhong et al., 2006). This approach has the advantage that it can detect novel dendritic mRNAs, but the likelihood of false‐ positives is high. Dendritic laminae in vivo contain the cell bodies of glial cells (astrocytes, oligodendrocytes, and resting microglia) as well as the cell bodies of various interneurons that reside in the laminae. Thus, it is important to verify the dendritic localization of candidate dendritic mRNAs identified in this way by in situ hybridization analyses, which has not yet been done for the novel candidates identified in this screen.

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Local Synthesis of Components of Multimolecular Signaling Complexes: Is There a Cotranslational Assembly?

It is noteworthy that several dendritic mRNAs encode proteins that are components of a highly organized multimolecular structure specialized for postsynaptic signal transduction termed the NMDA receptor complex (NRC) (Husi et al., 2000), including CAMKII, shank, the InsP3 receptor, and Arc. The existence of the complex was inferred by identifying the proteins that coprecipitate with the NMDA receptor (Husi et al., 2000). A related study that used mass spectroscopy to identify protein constituents of the ‘‘core postsynaptic density,’’ revealed a similar set of proteins (Walikonis et al., 2000) suggesting that the core postsynaptic density (psd) may be a scaffold made up largely of the NRC. Thus, the postsynaptic density/NRC appears to be a highly organized multimolecular structure specialized for postsynaptic signal transduction (Sheng and Lee, 2000). It is likely that proper signaling would require a precise stoichiometric relationship between the different molecules making up the NRC, raising the question of how such a complex is assembled. One possibility is that the complex is assembled away from the synapse, and inserted into the psd. Alternatively, local synthesis at synapses provides a mechanism that could allow the different molecular constituents of the NRC to be replaced by direct substitution into existing complexes at the postsynaptic density. In this regard, it is of interest that one of the components of the NRC (Arc) is expressed as an IEG, and disappears within hours after induction whereas the other component proteins have much longer half‐lives (Ehlers, 2003). If constituents of the NRC are replaced individually, then ribosomes and other components of the translational machinery would have to be closely associated, perhaps embedded within the postsynaptic density as they synthesize molecules of the NRC. Direct electron microscopic visualization of ribosomes is problematic because of the electron dense nature of the postsynaptic density, but studies using subcellular fractionation and electron microscopic immunocytochemical techniques have provided evidence for the localization of several components of the translational machinery and ribosomal protein in postsynaptic densities, which would be consistent with the presence of ribosomes in association with the density (Asaki et al., 2003). It is also noteworthy that strong synaptic activation, which is associated with an increase in the synthesis of molecules assembled into the NRC (Arc and CAMKII), causes a translocation of ribosomes from the spine base out into the spine head, where they would be closer to the postsynaptic density itself (Ostroff et al., 2002). Moreover, a more recent study reveals that a similar stimulation paradigm caused a threefold increase in the levels of CAMKII mRNA in isolated synaptodendrosomes without any change in overall CAMKII mRNA levels, suggesting translocation of preexisting CAMKII mRNA from the shaft of the dendrite out into the spine head (Havik et al., 2003). The translocation of polyribosomes and mRNAs encoding components of the NRC into the spine cytoplasm would mean newly synthesized proteins would emerge from the translational apparatus very close to the psd.


8

4

Dendritic Transport of RNA

Studies in cell types other than neurons have revealed features of mRNA trafficking mechanisms (Steward and Singer, 1997). Some of the key ideas are: (1) cells possess selective mRNA transport mechanisms that deliver appropriately addressed mRNAs to particular subcellular destinations; (2) mRNAs are selected for transport based on RNA transport signals within the mRNA (zip codes) (see Steward and Singer (1997)); mRNAs that do not contain transport signals remain localized in the perinuclear cytoplasm. (3) Certain mRNAs also contain address markers that cause the mRNA to localize selectively in particular subcellular domains.

8.1

Messenger RNA Transport Mechanisms

It is now clear that neurons use the same mRNA trafficking mechanisms. Early studies evaluated RNA transport mechanisms by pulse‐labeling hippocampal neurons in culture with 3H‐uridine, and following the movement of the newly synthesized RNA by assessing its distribution at different times after the pulse‐labeling period using autoradiography. These analyses revealed a dendrite‐specific RNA transport mechanism that delivered newly synthesized RNA at a rate of about 0.5 mm per day (Davis et al., 1987, 1990.). Given that about 80% of the label was in the rRNA, the bulk transport visualized by pulse labeling with uridine probably reflects the transport of ribosomes. More recent studies have revealed that mRNAs are transported at a much faster rate than suggested by the pulse‐labeling studies. For example, in situ hybridization analyses of the distribution of newly synthesized Arc mRNA after it has been induced by a seizure revealed that the mRNA is delivered throughout the dendrites of dentate granule cells by about 1 h (a distance of approximately 350 mm, so approximately 350 mm/h). Studies of movement of RNA‐containing granules in the dendrites of neurons in culture reveal rates that are remarkably similar (Knowles et al., 1996). More recent studies in which movement of Arc mRNA has been directly visualized in living cells reveal that exogenously expressed constructs made up of the 30 ‐UTR of Arc mRNA together with a reporter move at a variety of rates, the fastest of which (almost 70 mm per minute) is capable of delivering packets of Arc mRNA from the nucleus to distal dendrites in minutes (Dynes and Steward, 2007). At this rate, transport is almost certainly mediated by microtubules. The slower rates may be mediated by actin/myosin‐based transport mechanisms.

8.2

Cis‐Acting Sequences That Identify mRNA for Dendritic Transport

Because some mRNAs are abundant in dendrites whereas most are restricted to the region of the cell body, there must be some mechanism that achieves this sorting. Studies of mRNA sorting in oocytes, Drosophila embryos, and other spatially complex cell types (fibroblasts and oligodendrocytes) indicate that cis‐acting elements (usually in the 30 UTR) act as ‘‘zip codes’’ identifying mRNAs for transport away from the cell body (for a review, see (Steward and Singer (1997)). The same seems to apply to the delivery of mRNA into dendrites (Tiedge et al., 1999). The general location of putative dendritic targeting sequences have been identified for MAP2 (Blichenberg et al., 1999), CAMKII (Mayford et al., 1996; Rook et al., 2000), Arc (Kobayashi et al., 2005), and the noncoding pol‐3 transcript BC1 (Muslimov et al., 1997). In most cases, the evidence has come from studies in which fusion transcripts containing parts of the 30 ‐UTR have been introduced into neurons in culture. In the case of BC1, radioactive transcripts were synthesized and injected directly into neurons, and their distribution was assessed autoradiographically. In the other cases, fusion transcripts were expressed by transfecting cells with expression vectors, and the distribution of the fusion constructs was assessed by in situ hybridization. The critical measures were the number of cells exhibiting dendritic labeling and/or the distance of labeling along dendrites. Although these studies reported significant differences in the degree of

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dendritic labeling following transfections with reporter constructs containing critical parts of the 30 ‐UTR versus control constructs, the data were not completely satisfying. The reason is that significant dendritic labeling was also seen with control constructs that theoretically should not be delivered into dendrites. For example, in most studies, about 10% of the cells exhibit dendritic labeling with control transcripts whereas 35%–45% of the cells exhibit dendritic labeling with transcripts that contain the putative dendritic transport sequence. There is some concern that constructs injected into cells or that are expressed by vectors may not be sorted in the same way as native transcripts. Exogenously expressed transcripts are not processed in the nucleus in the same way as native transcripts, and transcripts expressed by vectors are often expressed at overly high levels, which could saturate the sorting machinery. Studies of mRNA transport for myelin basic protein (MBP) in oligodendrocytes have identified a number of considerations relating to the construction of fusion transcripts (Ainger et al., 1997). For example, it appears to be necessary that fusion transcripts be of a minimum size (larger than 500 bases). Also, although localization of MBP mRNA depends on a particular base sequence in the 30 ‐UTR, proper localization requires that the mRNA also contains an open reading frame. These considerations make it difficult to define the minimal sequences that are necessary and sufficient for dendritic transport. If exogenously expressed transcripts are sorted less accurately than transcripts transcribed by normal nuclear machinery, one would expect that the most accurate sorting assays would involve transcripts from transgenes introduced into mice. Indeed, one study did just this, and the differences in localization of transcripts containing the dendritic targeting sequence and control transcripts was striking. In this study, a transgenic mouse was produced in which in which most of the 30 ‐UTR of a‐CAMKII mRNA was replaced by the 30 ‐UTR of bovine growth hormone (Miller et al., 2002). In situ hybridization analyses revealed that CAMKII mRNA lacking the 30 ‐UTR remained in the cell body, and biochemical analyses revealed that CAMKII protein levels in the postsynaptic density were reduced. Physiological and behavioral studies revealed subtle but significant deficits in late‐phase LTP and memory in the mutant animals, suggesting that local synthesis of CAMKII protein is important for these processes. It cannot be excluded, however, that local synthesis is required to maintain high levels of a‐CAMKII protein in the psd, which in turn is important for the signal transduction events that are critical for late‐phase LTP. In this case, local synthesis itself would not play a direct role in bringing about the late‐phase modifications, but instead would create a signal transduction‐competent synapse (Steward, 2002a). Transgenic mice clearly provide a powerful assay for the existence and functional relevance of sequences involved in mRNA delivery. At the same time, transgenic approaches are certainly not the most efficient way to identify cis‐acting elements and will not be appropriate for testing many different constructs. Thus, less satisfactory approaches involving transfection of neurons in culture will almost certainly continue to be used.

8.3

Dendrite‐Specific RNA Transport

Various mRNAs are localized in the dendrites of mature neurons in vivo, but these same mRNAs are not detected in axons. Also, when neurons in culture are pulse‐labeled with uridine, newly synthesized RNA is transported throughout dendrites but not axons (Davis et al., 1987, 1990). The degree of selectivity is quite impressive; for example, when neurons in culture that has been pulse labeled with 3H uridine, and prepared for autoradiography 24 h later, dendrites are extensively labeled, whereas there is essentially no detectable labeling in axons. Thus, mRNA could either be excluded from the axon by some mechanism (a gate), or there must be something unique about the dendritic transport machinery that distinguishes it from the transport machinery in axons. In neurons and other spatially complex cells, transport over long distances occurs along tracks (generally microtubules) and involves different molecular motors, which carry specific cargoes. Selectivity is determined by the direction in which the motor molecule moves along polarized microtubules and upon interaction of the cargoes with the motor molecules. An extensive literature exists regarding mechanisms of transport of proteins and membrane vesicles; and in general, proteins of the kinesin family transport


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cargoes in an orthograde direction (away from the cell body) whereas dynein transports cargoes in the retrograde direction. Microtubules in axons are polarized (plus end distal) whereas microtubules in dendrites are of mixed polarity except in distal dendrites, where microtubule orientation is again primarily plus end distal (for a recent review see Hirokawa (2006)). There have been far fewer studies of molecules involved in RNA transport (Kanai et al., 2004; Hirokawa, 2006). One general conclusion about RNA transport in cells is that the cargo is not naked mRNA but is instead some sort of ‘‘RNA granule’’ made up of mRNA (sometimes multiple copies) and a variety of mRNA‐binding proteins, including proteins involved in translation (for a recent review of RNA granules, see Anderson and Kedersha (2006)). The same is true in neurons, and the composition of granules is an area of active investigation (Hirokawa, 2006). Building upon the core idea that mRNA is transported in granules, Hirokawa and colleagues developed a detergent extraction procedure to isolate large RNA‐containing granules associated with the conventional kinase KIF3, and then used proteomic approaches to identify proteins present in the granule (Kanai et al., 2004). Altogether, 42 different proteins were identified, and shown to be colocalized in individual granules in hippocampal neurons in culture. Although these studies have provided important information about the molecules in RNA granules, the critical motor (KIF3) also operates in axons. Thus, it is still not clear what differentiates the tracks in dendrites from those in axons, so as to account for the highly selective dendritic transport of RNA. One difference between dendritic and axonal microtubules is the complement of microtubule‐associated proteins (Binder et al., 1986), but whether and how these regulate the specificity of RNA transport is unknown.

8.4

Mechanisms Underlying mRNA Localization at Synapses

Studies of Arc mRNA have demonstrated that mechanisms exist to target the mRNA to active synapses. Even in the case of Arc mRNA that is already present throughout dendrites, synaptic activation triggers rapid relocalization to active synaptic sites within minutes (Steward et al., 1998b). Importantly, the relocalization is accompanied by decreases in mRNA in nonactivated dendritic segments. This implies that when synapses on middle dendritic regions are activated, Arc mRNA present in distal dendrites moves back toward the cell body; mRNA that is present in proximal segments moves distally, and then accumulates in the activated middle portion of the dendrite. In principle, relocalization of Arc mRNA could occur in atleast three ways: (1) there could be some vectorial transport toward the activated dendritic domain; (2) mRNAs could be moving continuously in orthograde and retrograde directions, and then dock selectively when an appropriate signal is generated by synaptic activation; (3) neuronal activation could destabilize mRNA in inactive dendritic segments and stabilize mRNA in the dendritic domains that were synaptically activated. It is hard to envision a transport mechanism that could result in vectorial transport toward an active synapse (especially when the vector of movement must be opposite in proximal versus distal dendritic segments). The third mechanism is also unlikely because the initial stages of relocalization occur without obvious decreases in mRNA levels in inactive segments. Accordingly, our working model is that Arc mRNA is being transported bidirectionally, and that synaptic activation creates some signal that causes mRNA to dock (or at least pause) in the activated dendritic domain. It is important to note, however, that direct evidence for such a docking mechanism is still lacking, and it cannot be excluded that activation of one dendritic domain triggers vectorial transport toward the active synaptic sites or that mRNA degradation/ stabilization contributes to the selectivity of localization. The nature of the docking mechanism, if it exists, remains to be defined.

9

Regulation of Translation of Dendritic mRNAs

It is now clear that a variety of different cell types, from oocytes to neurons, depend on transport of mRNA to particular intracellular destinations where the mRNAs are locally translated. If it is important to

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synthesize proteins in distinct subcellular compartments, logic suggests that it would also be important to repress translation of the localized mRNAs until they have reached their appropriate destination. Thus, it is assumed that translation of localized (i.e., dendritic) mRNAs is actively repressed during the time that the mRNA move from its site of synthesis in the nucleus to its final destination. A corollary concept is that mRNA transport and translational repression are linked. These concepts, and the evidence for them, come mostly from studies of maternal mRNAs that are selectively transported to different parts of oocytes or that are partitioned to different cell populations during early embryogenesis.

9.1

Translational Repression of Localized mRNAs in Neurons

In neurons, the most well‐developed story pertains to the mRNA for b‐actin, and its repression by a protein that binds to the portion of the mRNA that serves as a ‘‘zip code’’ for transport (the zip code‐binding protein ZBP1). The mRNA for b‐actin is one of the mRNAs that is transported into axonal growth comes of young neurons (Bassell et al., 1998). It is thought that local synthesis of b‐actin plays a critical role in regulating cytoskeletal dynamics during growth cone motility, and in creating the local protrusions that are critical for growth cone motility and extension (Bassell, 1998). It has been shown that the delivery of b‐actin mRNA into axons depends on a sequence in the 30 ‐UTR that functions as a ‘‘zip code.’’ The presence of this sequence is necessary and sufficient to cause reporter mRNA constructs containing the sequence to be transported into axons. It has been established that mRNA transport and localization are mediated by mRNA‐binding proteins, and a search for proteins that bind the zip code sequence identified a protein, now termed ZBP1, that binds the appropriate portion of the 30 ‐UTR and was colocalized with b‐actin mRNA during transport (Tiruchinapalli et al., 2003). Subsequent studies then revealed that ZBP1 plays a role in both mRNA transport and translational repression, binding the mRNA while it is still in the nucleus, and then repressing translation initiation once the mRNA reaches the cytoplasm (Huttelmaier et al., 2005). This translation block is then released when the mRNA reaches its final destination in the growth cone, and subsequent translation is then regulated by other signal transduction mechanisms.

9.2

Translational Repression of Dendritic mRNAs While They Are in Transit

Although the concept of translational repression of localized mRNA is compelling on first principles, direct evidence for translational repression of known dendritic mRNAs is sparse. The best evidence pertains to the mRNA for CAMKII and involves a mechanism that was first elucidated in studies of repression of maternal mRNAs in oocytes (Wells, 2006). Many maternal mRNAs present in oocytes are translationally repressed until fertilization and have truncated poly(A) tails. Translational repression is due to the presence of a sequence in the 30 ‐UTR (a cytoplasmic polyadenylation element or CPE) that binds a protein called cytoplasmic polyadenylation element‐binding protein (CPEB). CPEB interacts with a 4E‐binding protein called maskin, which forms a complex with the 50 ‐cap that prevents the formation of the initiation complex. Upon fertilization, CPEB is phosphorylated by a specific kinase called aurora kinase, which causes CPEB to dissociate from maskin, which then results in the recruitment of a complement of proteins that extend the poly(A) tail of the mRNA and recruits the initiation complex triggering translation. Neurons also express CPEB, and the protein is present in dendrites and localized at synapses. In neurons, CPEB can be phosphorylated by either aurora kinase or CAMKII, which is of interest because CAMKII is one of the signaling molecules regulated by synaptic (especially NMDA receptor) activation. Of the known dendritic mRNAs, only the mRNA for CAMKII contains a consensus CPE, which is a target for CPEB, and there is direct evidence for cytoplasmic polyadenylation of CAMKII mRNA and translational activation in response to physiological signals (Wu et al., 1998; Wells et al., 2001; Huang et al., 2002). Interestingly, this mechanism could be triggered by behavioral experience (the first light exposure


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for animals that had been raised in the dark). Subsequent immunocytochemical studies revealed that a number of factors known to control polyadenylation‐induced translation in oocytes were present in dendrites, and regulated by NMDA receptor‐dependent signal transduction pathways (Huang et al., 2002). Moreover, glutamate stimulation of neurons in culture triggered translation of reporter constructs that contained intact CPEs (Wells et al., 2001). Together, these results document the presence of a mechanism capable of regulating local translation of CAMKII mRNA at synapses in response to signals generated by synaptic activation. Although this mechanism appears to be important for regulating the translation of CAMKII mRNA, it is noteworthy that CPEs have not been identified in other known dendritic mRNAs. The mechanism of translational repression involving CPEB involves several proteins, and it is of interest that the mechanism of repression would create a circular complex of the mRNA and associated proteins (Wells, 2006). This again raises the issue of spatial constraints in the cytoplasmic microenvironment of the spine and especially the spine neck. The individual proteins are relatively small in relation to the total complex, but a single copy of CAMKII mRNA with all these proteins attached would be relatively large in comparison to the neck of a typical spine, which could limit the ability of the CAMKII mRNA complex to move into the spine. Yet there is evidence that the mRNA for CAMKII actually does move into spines in response to the induction of LTP (Havik et al., 2003). If CPEB and maskin are important for repressing translation as the mRNA is in transit, these proteins may dissociate completely from the mRNA, which would reduce the overall size of the complex, but there are still a number of proteins that would presumably be bound. Alternatively, it is conceivable that spines are like snakes that can swallow things larger than their maximal diameter, expanding to accommodate the mass.

9.3

The Fragile‐X/BC1 Connection

Still another mechanism for controlling translation at synapses involves Fragile X mental retardation protein (FMRP). Fragile X mental retardation syndrome is caused by a mutation in the gene encoding FMRP (usually an expanded trinucleotide repeat that is hypermethylated, inhibiting gene transcription). FMRP is an RNA ‐binding protein, and electron microscopic immunocytochemical studies revealed that FMR protein is concentrated around SPRCs (Feng et al., 1997). On this basis, Feng and coworkers proposed that FMRP might be involved in targeting mRNAs to dendrites or regulating their translation. Studies of FMRP‐knockout mice revealed that there were no gross abnormalities in the dendritic localization of representative dendritic mRNAs (MAP2, CAMKII, and Arc), but this study did not exclude the possibility of subtle deficits in mRNA targeting (Steward et al., 1998a). Subsequent studies sought to define the mRNAs bound by FMRP. One identified a ‘‘G quartet’’ domain, which appears to be one motif that mediates binding of mRNA to FMRP, but none of the principle ‘‘dendritic’’ mRNAs (> Table 4-1) have this domain (Darnell et al., 2001). Another study identified a different set of mRNAs that interact with FMRP (Miyashiro et al., 2003), some of which are localized in proximal dendrites, but again none of the mRNAs that are abundant in dendrites turned up. A new twist to the story has come from a recent study that indicates that FMRP acts as a repressor of translation of several of the principle dendritic mRNAs including CAMKII, and Arc, as well as b‐actin and FXR2 (a protein related to FMRP) (Zalfa et al., 2003). This study also showed that FMRP interacts with the regulated mRNAs via a noncoding pol‐3 transcript called BC1, which has previously been shown to be localized in dendrites (Tiedge et al., 1991). Interestingly, BC1 contains sequences that are predicted to base pair with sequences in MAP1B, CAMKII, and Arc mRNAs. These results suggest that BC1 may link particular mRNAs to FMRP, leading to repression of translation. This ties in nicely with other work implicating BC1 as a regulator of translation initiation of dendritic mRNAs (Wang et al., 2002). These findings have led to the interesting idea that the loss of FMRP in fragile X mental retardation syndrome could lead to a dysregulation of mRNA translation at the synapse, disrupting synaptic function (Zalfa et al., 2003).

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Micro‐RNAs

Recent studies have raised the possibility that micro‐RNAs play a role in regulating the translation of dendritic mRNAs. For example, the micro‐RNA miR‐134 is abundant in dendrites of young neurons in culture, where it seems to play a role in regulating the translation of another dendritic mRNA (Limk1) that is present in the dendrites of developing neurons (Schratt et al., 2006). It is not yet known whether the micro‐RNAs present in the dendrites of neurons in culture are also present in adult dendrites in vivo. This is clearly a rapidly emerging area of investigation, and it is likely that this story will develop rapidly over the coming years.

9.5

Regulation of Translation by Synaptic Signals

The selective localization of polyribosomes at synapses invites the speculation that signals generated by synaptic activity may regulate translation of mRNAs in dendrites. As discussed above, however, most synapses have only one or two polyribosome clusters (Steward, 1983). If each cluster is associated with an individual mRNA, this means that one or two mRNAs are being translated at any given time. Given that there are a number of different mRNAs present in dendrites (even with the more conservative list of mRNAs that are evident by in situ hybridization), there must be competition for initiation and translation. How this is orchestrated is just now being investigated, and it appears that the story will be complex. Studies of the local translation of CAMKII mRNA have suggested that local synthesis is regulated via NMDA receptor activation. In two studies, it was shown that high‐frequency stimulation designed to induce LTP (which activates NMDA receptors) led to rapid increase in immunostaining for CAMKII within dendrites. One study involved hippocampal neurons in slices (Ouyang et al., 1999); the other involved dentate granule neurons in vivo (Steward and Halpain, 1999). The increases in immunostaining were blocked by NMDA receptor antagonists (Steward and Halpain, 1999), implying a role for NMDA receptors in translational activation. One puzzle was that increases in immunostaining were blocked by protein synthesis inhibitors applied to hippocampal slices (Ouyang et al., 1999), but not when inhibitors were delivered in vivo (Steward and Halpain, 1999). Subsequent studies of CAMKII synthesis in isolated synaptodendrosomes provided clarification on the issue of sensitivity to protein synthesis inhibitors, and also on the mechanisms of translational activation (Sheetz et al., 2000). CAMKII synthesis in synaptodendrosomes (as measured by incorporation of 35S methionine into CAMKII protein) was enhanced by NMDA receptor activation. Interestingly, there was also increased phosphorylation of the initiation factor eIF2, which would be expected to decrease the rate of polypeptide elongation. This apparent paradox can be explained by the fact that decreases in elongation rate favor the translation of weakly initiated mRNAs, and CAMKII is one of the mRNAs for which initiation is inefficient. Consequently, decreases in elongation consequent to eIF2 phosphorylation could lead to increases in CAMKII synthesis. In support of this idea, Sheetz and coworkers showed that low to moderate concentrations of cycloheximide (which inhibit elongation) increased incorporation into CAMKII at the same time that overall levels of protein synthesis were diminished. These results may explain why protein synthesis inhibitors failed to block the increases in immunostaining after synaptic activation in vivo (Steward and Halpain, 1999). It remains to be seen how these mechanisms interact with the mechanism involving CPEB, described above. Whereas the evidence to date indicates that translation of CAMKII is regulated via NMDA receptor activation, translation of other mRNAs may be regulated in other ways. Local synthesis of FMRP appears to be regulated by mGluR activation; for example, treatment of synaptoneurosomes with agonists for metabotropic glutamate receptors causes a rapid increase in the amount of FMRP detectable by Western blotting (Weiler et al., 1997). The translation of other dendritic mRNAs appears to be insensitive to neurotransmitter activation. For example, using another measure of translation (association of mRNAs with polysomes) (Bagni et al., 2000), it was confirmed that glutamate application or depolarization recruited CAMKII mRNA to polyribosomes, but did not recruit mRNAs for InsP3R1 or Arc.


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Another recent study reports, however, that Arc mRNA translation in synaptosomes is upregulated by exogenous recombinant reelin acting through integrin receptors (Dong et al., 2003). This induction was blocked by echistatin (which blocks integrin receptors) and by rapamycin, implying that translation is regulated through the rapamycin‐sensitive kinase mammalian‐target‐of‐rapamycin (mTOR, a.k.a. FRAP kinase and RAFT‐1). Previous studies had shown that several putative components of this translational signaling pathway, including mTOR, 4E‐BP1, 4E‐BP‐2, and eIF‐4E are present in dendrites (Tang et al., 1998), and rapamycin blocks several forms of protein synthesis‐dependent synaptic plasticity (Tang et al., 1998; Steward and Schuman, 2001). It remains to be seen how integrin receptor‐mediated signals and signals generated by neurotransmitters are integrated by the translational machinery at synapses. Adding still further to the complexity is the evidence that several dendritically localized mRNAs have internal ribosome entry sites (IRESs), including CAMKII, Arc, dendrin, MAP2, and RC3, and that these mRNAs can be translated in a cap‐dependent or cap‐independent fashion (Pinkstaff et al., 2001). Interestingly, studies of bi‐cistronic constructs with two different reporters revealed that IRES‐mediated translation was relatively more efficient in dendrites (Pinkstaff et al., 2001), and studies using bi‐cistronic constructs in Aplysia neurons have revealed that egg‐laying hormone, which triggers a bout of intense activity, causes a switch from cap‐dependent to cap‐independent translation (Dyer et al., 2003). It remains to be seen whether IRES‐mediated translation is regulated by synaptic activation in mammalian neurons or by other signals impinging on dendrites.

9.6

Regulation of Protein Synthesis in Developing Dendrites by Miniature Synaptic Events

A new twist to the story of synaptic regulation is the evidence that miniature synaptic events (mini’s) regulate translation of dendritic mRNAs in developing dendrites differently than the synaptic events triggered by presynaptic action potentials. Using a GFP‐based reporter system, Sutton and coworkers showed that dendritic protein synthesis was modestly affected by blockade of action potential‐dependent transmission, and disruption of miniature synaptic transmission led to dramatic increases in local protein synthesis (Sutton et al., 2004) and surface synaptic GluR1 receptors (Sutton et al., 2006). Moreover, the protein synthesis‐dependent increase in GluR1 leads to the insertion of a novel, Ca2þ‐permeable GluR channel. In a related study, prolonged (3 days) blockade of activity (using TTX and APV) was reported to increase the levels of a transfected glutamate receptor in the dendrites (Ju et al., 2004). These results suggest the hypothesis that minis may serve as a signal for synaptic integrity at synapses; the presence of minis keeps the protein synthesis machinery in a repressed state, whereas a loss of minis results in a stimulation of protein synthesis, which triggers an increase in GluR insertion in order to compensate for net decreases in presynaptic input. The story regarding translational regulation of dendritic mRNAs is rapidly evolving, and the final answer is likely to be complex, especially given the limited protein synthetic machinery at individual synapses. One possibility is that enhancement of translation of one type of mRNA would lead to decreased translation of other mRNAs because of competition, but this remains to be explored. Many questions remain about how all this is regulated by signals generated by particular neurotransmitter receptors.

10

Protein Synthesis in Axons

This chapter has focused on protein synthesis in dendrites, but we note that local protein synthesis also occurs in axons (for a review, see Giuditta et al. (2002)). Messenger RNAs and translational machinery are present in the neurites of invertebrates, which have the characteristics of both axons and dendrites (van Minnen and Syed, 2001). Studies of Aplysia have revealed that neuritic protein synthesis is critical for several different forms of activity‐dependent synaptic plasticity (Martin et al., 1997; Sutton et al., 2001). It is

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not yet clear whether the critical protein synthesis‐dependent events are on the presynaptic or postsynaptic side, or whether different mechanisms operate for different types of synaptic modification. In adult vertebrate axons, polyribosomes are present beneath synapses on axon initial segments (Steward and Ribak, 1986) but are generally not detectable in distal axons. Messenger RNAs encoding the neuropeptide neurotransmitters oxytocin, vasopressin, and pro‐dynorphin are present in axon terminals of the hypothalamo‐hypophyseal tract (Mohr et al., 1991), and mRNAs for the olfactory marker protein and various odorant receptors are present in the axon terminals of olfactory neurons that terminate in the olfactory bulb (Ressler et al., 1994; Vassar et al., 1994). The significance of the localization of mRNAs in these axons is not clear, because ribosomes have not been detected in these axon terminals.

10.1 Protein Synthesis in Growing Axons and Growth Cones Local protein synthesis appears to be especially important in growing axons, especially within growth cones. Local synthesis of b‐actin is important for cytoskeletal remodeling at the leading edge of the growth cone and within filopodia (Bassell et al., 1998), and signals that stimulate axon growth also trigger increases in the levels of b‐actin mRNA in growth cones (Zhang et al., 1999). As noted above, localization of b‐actin mRNA appears to be mediated by a cis‐acting sequence that is recognized by zip code‐binding protein (ZBP). Disruption of the interaction between ZBP and b‐actin mRNA disrupted mRNA localization, reduced b‐actin protein levels within growth cones, and impaired growth cone motility (Zhang et al., 2001). In contrast to the situation in adult organisms, where polyribosomes are not detected in axons except in initial segments, polyribosomes are abundant in growth cones of at least some growing axons in vitro (Bassell et al., 1998) and in vivo (Tennyson, 1970). It is interesting that the significance of the latter observation is only now becoming apparent. Recent evidence also implicates local protein synthesis within growth cones in growth cone turning in response to guidance cues. Growth cones extend and retract filopodia and ruffled membranes and the net direction of extension is determined by where extension and retraction occur. Turning is caused by extension on one side and collapse on the other and is triggered by local attractive and repulsive cues in the environment. Studies of chemotropic responses of growth cones of Xenopus retinal ganglion cells have demonstrated that growth cone collapse and turning away in response to sema3A are blocked by protein synthesis inhibitors, even in growth cones that had been separated from their cell bodies (Campbell and Holt, 2001). Protein synthesis inhibitors also blocked the attractive turning response normally seen when retinal growth cones from young embryos were exposed to a gradient of netrin‐1, and also the repulsive turning induced by netrin‐1 when neurons were grown on laminin. Together, these results indicated that local protein synthesis within the growth cone is essential for both repulsive and attractive guidance mechanisms. Moreover, exposure to sema3A or netrin‐1 triggered a burst of protein synthesis within growth cones. One paradox is that protein synthesis is important both for collapse (a deconstruction of the growth cone) and for turning toward netrin‐1 (a positive response). A role for local protein synthesis in growth toward an attractant is consistent with the idea that local synthesis is important for extension of the growth cone. What role new protein synthesis plays in growth cone collapse remains to be established, but the opposite responses to netrin‐1 indicate that particular extracellular signals can be ‘‘translated’’ in different ways. A somewhat different role for local protein synthesis was suggested by studies that revealed adaptation of Xenopus spinal growth cones to the chemoattraction produced by gradients of netrin‐1 and BDNF (Ming et al., 2002). The main finding was that attractive turning of the growth cone toward a netrin‐1 or BDNF source was attenuated by the presence of background concentrations of either chemoattractant in the bath. This ‘‘de‐sensitization’’ disappeared within 30 min after removing netrin‐1 from the bath, and resensitization was blocked by protein synthesis inhibitors. A still different role for local protein synthesis in axonal growth cones is suggested by the work of Brittis and coworkers (2002), who documented the existence of a mechanism that could allow the local synthesis


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of membrane receptors for axon guidance molecules. Spinal commissural axons grow initially toward the midline floor plate in response to attractive guidance cues. After crossing the midline, the axons lose their responsiveness to midline attractants and gain responsiveness to a new set of guidance cues so that the axons grow longitudinally toward the brain (Stein and Tessier‐Lavigne, 2001). One possible explanation for the change in responsiveness is a local synthesis of receptors for guidance molecules just after growing commissural axons reached the midline. Evidence for such a mechanism came from studies of the expression of an EphA2 receptor–GFP reporter construct in commissural axons. EphA2 is one of several proteins that are expressed selectively on the distal segments of axons of commissural neurons after they have crossed the midline. Commissural neurons transfected with constructs made up of the 30 ‐UTR of EphA2 and a fluorescent reporter protein (GFP) exhibited protein expression in cell bodies, but not in proximal segments of the axons prior to midline crossing. In contrast, the protein was expressed at high levels in the distal segments of the axons that had extended beyond the midline. These results reveal a mechanism that could allow the local synthesis of receptors for guidance cues. One can imagine a scenario in which one set of receptors are expressed until growing axons reach intermediate point A; signals from point A then trigger the translation of mRNAs for a different set of receptors critical to guiding the axon to the next intermediate station. It should be emphasized, however, that the study involved an exogenously transfected reporter that had the 30 ‐UTR from EphA receptor mRNA, and did not directly demonstrate the presence of EphA receptor mRNA in growing axons. It also remains to be established whether there is functional ER and Golgi apparatus in axonal growth cones. A key proof of principle would be to show that disrupting local synthesis would disrupt axon guidance. Interesting new evidence indicates that axonal transport of mRNA may be reinitiated during axonal regeneration. Regenerating axons of adult dorsal root ganglion cells and spinal motoneurons contain ribosomal proteins, translation initiation factors, and rRNA (Zheng et al., 2001). Axons of dorsal root ganglion cells that have been induced to regenerate by a conditioning lesion also contain mRNAs for action and neurofilament protein, and blocking protein synthesis within these axons causes growth cone retraction. Together, these findings suggest that local protein synthesis is a critical factor in successful axon regeneration.

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Degradation of Dendritic mRNAs and Locally Synthesized Proteins

Local protein synthesis at synapses has great appeal as a mechanism to regulate levels of particular proteins, and if this is valuable to the cell, cellular processes that allow local degradation of RNA and proteins should also be advantageous. The extent to which these processes can occur locally in dendrites remains to be established. Little is known about the half‐life of mRNA in dendrites (except for Arc mRNA) or the mechanism or site of mRNA degradation. P‐bodies, which may be an important site of mRNA degradation, appear to be mainly localized in the neuronal cell body, but it is not clear whether mRNA degradation takes place in locations that lack P‐bodies, and if so, through what mechanism. This is a very active field of investigation, and it is likely that the story will change rapidly over the next few years. Similarly, the mechanisms of protein degradation in dendrites are not yet fully known. The canonical mechanism for protein degradation is the ubiquitin‐proteasome system, in which proteins are marked for degradation by the addition of a ubiquitin chain, and then delivered to the 26S proteasome, where degradation occurs. The machinery required to carry out ubiquitin‐dependent proteolysis includes the ubiquitin‐conjugating enzymes (E1, E2, E3), ubiquitin, and the 26S proteasome, which is formed by the co‐ assembly of a 20S proteasome (the catalytic component) and a 19S cap (the regulatory component). Immunocytochemical studies have revealed that ubiquitin and the subunits of the proteasome are present in dendrites (Patrick et al., 2003), and ubiquitinated proteins have been detected in synaptic fractions from adult rat brains (Chapman et al., 1994). It has been shown that inhibition of proteasome activity blocks the agonist‐mediated internalization of both GluR1 and GluR2 in hippocampal neurons, revealing a mechanism by which the ubiquitin‐proteasome pathway can modulate synaptic transmission (Patrick et al., 2003).

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Information is only now beginning to emerge about how protein degradation is regulated by synaptic signals (Ehlers, 2003).

Acknowledgments The author’s research is supported by NIH‐NS12333 and by FRAXA.

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5

Emerging Functions of the ‘‘Ca2+ Buffers’’ Parvalbumin, Calbindin D-28k and Calretinin in the Brain

B. Schwaller

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198

2

Buffering vs. Sensor Function of CaBPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199

3 3.1 3.2 3.3 3.4 3.5

Important Parameters Describing the Properties of Ca2þ Buffers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 200 Intracellular Concentration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 200 Metal‐Binding Affinities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 Metal‐Binding Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204 Mobility and Interaction with Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204 Models Used to Investigate the Role of Synthetic Ca2þ Buffers and CaBPs . . . . . . . . . . . . . . . . . . . . . . 204

4

Distribution of Parvalbumin (PV), Calbindin D-28k (CB), and Calretinin (CR) in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 4.1 PV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 4.2 CB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 206 4.3 CR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 5 5.1 5.2 5.3 5.4

Transgenic Models Revealing the Functions of CaBPs in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208 PV Knockout Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208 CB Knockout‐ and CB Antisense Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209 CR Knockout Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 Multitransgenic and Reporter Strains: CaMII‐PV, Thy‐1‐PV, PV‐EGFP, and Cre‐PV . . . . . . . . . . . . 212

6 6.1 6.2 6.3

Alterations in CaBP Expression and Relation to Brain Pathologies . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212 Neuronal Loss vs. Loss of CaBP Immunoreactivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213 Correlation vs. Cause vs. Secondary Adaptive Changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213 The ‘‘Ca2þ Homeostasome’’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214

7

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215

#


198

5

Emerging functions of the ‘‘Ca2+ buffers’’ parvalbumin, calbindin D-28k and calretinin in the brain

Abstract: Together with the ubiquitous calmodulin (CaM), the EF‐hand containing calcium‐binding proteins (CaBPs), parvalbumin (PV), calbindin D‐28k (CB), and calretinin (CR), are the most abundantly expressed members of this family in the brain. Formerly, they were classified as simple buffers serving to ‘‘clamp’’ the intracellular calcium concentration [Ca2þ]i. But recent studies often using transgenic mice have revealed these molecules to play pivotal roles in Ca2þ homeostasis and signaling. And research conducted during the last 5 years indicates that they are important for synaptic plasticity and related rhythmic activities within neuronal networks. For CB, an additional modulator role in inositol‐1,4,5‐ trisphosphate (IP3)‐signaling pathways was reported, indicating additional sensor functions. In this chapter, I summarize the current knowledge on the three CaBPs in the brain revealing their important roles in the CNS. List of Abbreviations: AHS, Ammon’s horn sclerosis; ALS, amyotrophic lateral sclerosis; BAC, bacterial chromosome; Ca2+, calcium; CaM, calmodulin; CABPs, calcium-binding proteins; CB, calbindin D-28k; CCK, cholecytokinin; CICR, Ca2+-induced Ca2+ release; CNS, central nervous system; CR, calretinin; FRAP, fluorescence recovery after photobleaching; EGFP, enhanced green fluorescent protein; GAD67, glutamate decarboxylase; IP3, inositol-1,4,5-triphosphate; IPSP, inhibitory postsynaptic potential; LTD, long-term depression; LTP, long-term potentiation; MNTB, medial nucleus of the trapezoid body; NCS, neuronal calcium sensors; NCXs, Na+/Ca2+ exchangers; PMCAs, plasmalemmal Ca2+-ATPases; PTZ, pentylenetetrazole; PV, parvalbumin; RTN, thalamic reticular nucleus; SERCAs, sarcoplasmic reticulum Ca2+-ATPases

1

Introduction

Calcium ions (Ca2þ) serve as an indispensable second messenger in many vital cellular functions. Processes as diverse as fertilization, muscle contraction, neuronal signaling, cell cycle regulation, and regulated cell death depend on spatially and temporally precise Ca2þ signals (Berridge, 1998; Berridge et al., 2003). In neurons, Ca2þ signals play key roles in intrinsic membrane excitability, gene transcription, neurotransmitter release and associated synaptic plasticity (long‐term potentiation (LTP) and long‐term depression (LTD)), oscillatory activity, and neuronal motility and morphology, including the growth and branching of dendrites and the formation of dendritic spines. To achieve precision in Ca2þ signaling, cells express specific components, which permit the influx of this cation across the plasma membrane or its release from intracellular stores such as the endoplasmic reticulum or mitochondria. A Ca2þ transient is terminated by the activity of specific Ca2þ pumps, which extrude the cation either into the extracellular space or back into the intracellular stores. Plasmalemmal Ca2þ‐ATPases (PMCAs) and Naþ/Ca2þ exchangers (NCXs) are involved in the former process, while the sarcoplasmic reticulum Ca2þ‐ATPases (SERCAs) and the yet uncharacterized mitochondrial Ca2þ uniporter are responsible for the latter. Cytosolic proteins capable of binding Ca2þ with precisely tuned affinities are important modulators of intracellular Ca2þ signals. Numerous possibilities exist by which Ca2þ‐binding sites in proteins may be constructed; several distinct protein families contain various evolutionarily well‐conserved Ca2þ‐binding domains. These include the EF‐hand proteins, the annexins, and the C2 domain proteins (for review see Celio et al., (1996); Schwaller, (2004)). Among the calcium‐binding proteins (CaBPs), those possessing the EF‐hand domain (Celio et al., 1996; Kawasaki et al., 1998) are the most common. Analysis of the human genome has revealed 242 such proteins (Lander et al., 2001). This feature also renders them one of the largest groups of proteins sharing a common motif. The canonical EF‐hand domain (helix–loop–helix motif) consists of a conserved stretch of usually 29 amino acids (two more in the S100‐specific N‐terminal Ca2þ‐binding loop), which are formed sequentially into an a helix, a Ca2þ‐binding loop, and a second a helix orientated perpendicular to the first (> Figure 5-1). The structure of this motif was elucidated by X‐ray analysis of carp parvalbumin (Kretsinger et al., 1973). The three helical pairs were named AB, CD, and EF, and the domain containing the C‐terminal a helices 5 and 6 (EF) conferred the entire family its name. The EF‐hand domain can be visualized by the thumb and the index of the right hand, which represent the two helices, and by the bent middle finger, which represents the loop providing the usually six oxygen atoms that coordinate Ca2þ. The seventh ligand is typically an oxygen atom from a water molecule. Hence, the seven oxygen ligands form a


5

. Figure 5-1 The EF‐hand motif. (a) The three‐dimensional organization of the EF‐hand motif can be simulated using the right hand: the index finger represents the E‐helix (residues 1–10), the bent middle finger symbolizes the 12 amino acids of the Ca2þ‐binding loop (10–21), and the thumb depicts the F‐helix (19–29). The seven oxygen ligands coordinating the Ca2þ ion are located in the seven corners of a pentagonal bipyramid (modified from Celio et al. (1996)). (b) X‐ray crystal structure from the EF‐domain of parvalbumin (modified from Kretsinger et al. (1973)). (c) Coordination of the Ca2þ ion in calmodulin (CaM) with the seven oxygen ligands: five from side chains, one from a carbonyl group of the backbone, and the seventh from a water molecule

pentagonal bipyramid (> Figure 5-1). An EF‐hand protein contains multiple, and usually an even number (two, four, six, or eight) of EF‐hand domains. The exceptions are parvalbumin and oncomodulin with three domains, and the family of penta‐EF‐hand proteins with five domains (Maki et al., 2002). Typically, not all EF‐hand domains are functional. For example, in parvalbumin only the CD and EF domains, not the AB site, bind Ca2þ. In this chapter, emphasis is placed on the CaBPs parvalbumin (PV), calbindin D‐28k (CB), and calretinin (CR), which represent the major species in the central nervous system (CNS).

2

Buffering vs. Sensor Function of CaBPs

The EF‐hand CaBPs are structurally and functionally grouped into two categories: Ca2þ sensors/modulators and Ca2þ buffers (Skelton et al., 1994; Ikura, 1996; Nelson et al., 1998) (see the chapter XY by Heizmann et al. (Leclerc et al., 2006)). Ca2þ sensors are characterized by their ability to undergo significant Ca2þ‐dependent conformational changes, which permit their interaction with specific targets in a Ca2þ‐ regulated manner. The archetypical, best‐described, and most representative example of the sensor class of EF‐hand proteins is calmodulin (CaM) (Cohen et al., 1988). It is a small protein (Mr 17 kDa; 148 amino acid residues) ubiquitously expressed and highly conserved among different species. In the presence of Ca2þ, the protein undergoes a conformational change from a dumbbell shape, with pairs of EF‐hand domains representing the two peripheral globes and a central long helix, to a more compact globular structure. CaM is implicated in many CNS functions, including synaptic transmission, neuronal plasticity associated with short‐term and long‐term potentiation, and learning and memory processes (see chapter XY by Heizmann et al. (Leclerc et al., 2006)) . Other families of Ca2þ of sensors include the S100 proteins (Marenholz et al., 2004) and the neuronal calcium sensors (NCS) (Braunewell et al., 1999; Burgoyne et al., 2001).

199

200

5


The two ‘‘classical’’ examples of Ca2þ‐buffering EF‐hand proteins are PV (official human gene symbol: PVALB) and calbindin D‐9k (also known as calbindin 3, CaBP‐9k, and CABP1; official gene symbol: S100G) (Marenholz et al., 2004). The latter is a member of the S100 family. Both proteins undergo conformational changes upon Ca2þ‐binding. But globally, the structure is essentially the same in the metal‐bound and free (apo) states (Skelton et al., 1995). To date, no binding partners for the two proteins have been reported, indicating their principally Ca2þ‐buffering roles. Among the species so far investigated, CB D‐9k has not been detected in the CNS. With respect to CR (also known as calbindin 2, protein 10 (Winsky et al., 1989) and calbindin 30 kDa (Hubbard et al., 1995); official gene symbol: CALB2) and CB D‐28k (also known as calbindin 1, CaBP‐28k, and spot 35 (Yamakuni et al., 1987); official gene symbol: CALB1) the situation is more complicated. In many reports, they are still considered as Ca2þ‐buffering proteins, but recent progress indicates that they have additional ‘‘sensor’’ functions.

3

Important Parameters Describing the Properties of Ca2þ Buffers

In many respects, the term ‘‘Ca2þ buffer’’ is rather a feeble one. But for mainly historical reasons and in the absence of a better alternative, it will still be used in this chapter. In chemical terms, a pH‐buffering system consists of an acid and base, which is optimized to ‘‘clamp’’ the pH to a predetermined value in such a way that the addition of an acid or a base elicits only a negligible change in the pH of the solution. Hence, the ‘‘buffering’’ capacity is highest when the pH is identical, or close to, the pK value of the corresponding acid– base pair. But this is not the case for Ca2þ‐buffering proteins. Under resting conditions, the intracellular cytosolic concentration of Ca2þ, [Ca2þ]i, is in almost all cells, including those of the CNS (neurons and glia), in the order of 50–100 nM. However the dissociation constants (Kd) of most CaBPs are in the low micromolar range. Thus, under resting conditions, most of the Ca2þ‐buffering protein molecules are in the Ca2þ‐free form. Nevertheless, if [Ca2þ]i increases due to Ca2þ influx through Ca2þ‐permeable channels or receptor channel complexes in the plasma membrane or by a release from internal stores, Ca2þ‐buffering proteins will modulate the spatiotemporal aspects of Ca2þ signals. The most important parameters relating to this buffering function include: the cytosolic concentration of the buffers (Sect. 2.1), their affinity for Ca2þ and other metal ions (Sect. 2.2), the kinetics of Ca2þ binding and release (Sect. 2.3), and their mobility (Sect. 2.4). These parameters have not been determined for PV, CB, and CR with precision in vivo. But recent progress in the field is summarized.

3.1 Intracellular Concentration An accurate determination of the cytosolic concentrations of CaBPs in neurons is not easy. Reported results were obtained using biochemical, immunocytochemical, radioimmunological (RIA), and electrophysiological methods. Additionally, the recording of [Ca2þ]i transients using fluorescent Ca2þ indicator dyes in combination with modeling the effects of buffers on [Ca2þ]i transients allows to estimate the binding properties and intracellular concentration. Evidently, in the latter case, the quality of the results is critically dependent on the quality of the metal‐binding parameters determined by various other methods (for methodological details, see Cox, 1996; Lee et al., 2000; Nagerl et al., 2000b). Each of the three proteins PV, CB, and CR is characterized by a very restricted pattern of expression within neurons of the CNS (see > Sect. 3), which, together with the complexity of neuronal morphologies, renders the determination of CaBP concentrations extremely difficult. In single, tall saccular hair cells of the frog, levels of CR— determined by Western blot—are extremely high (1.2 mM (Edmonds et al., 2000)), whereas in the inner and outer hair cells of the rat cochlea they are significantly lower. Quantitative immunohistochemistry and ultrastructural immunostaining have revealed these to be 19 2 mM and 35 3 mM, respectively (Hackney et al., 2005). A similar CR concentration range (30–40 mM) has been proposed for cerebellar granule cells; CR/ neurons display an altered neuronal excitability that can be compensated by injecting 150 mM BAPTA (Gall et al., 2003). Taking into account that CR has five Ca2þ‐binding sites (Cheung et al., 1993; Schwaller et al., 1997), 30 mM CR is roughly equivalent to 150 mM BAPTA, although this needs to be


5

considered with precaution (see Sect. 2.5). The CB‐concentration range in different neuronal populations is generally higher: 150–360 mM in cerebellar Purkinje cells (for review see Schwaller et al., (2002)), and 40–50 mM in mature hippocampal dentate gyrus granule cells, CA3 stratum radiatum interneurons, and in CA1 pyramidal cells as determined by quantitative immunohistochemistry (Muller et al., 2005). Estimates of the PV concentration in Purkinje cells range from 80 mM in the mouse (Schmidt et al., 2003b) to 116 30 mM in the rat (Hackney et al., 2005). A value of 150 mM for molecular‐layer interneurons of the murine cerebellum fits best with the experimental data (Collin et al., 2005). Using ELISA and ultrastructural immunostaining methods, values ranging from 50 mM (Plogmann et al., 1993) up to 1 mM (Kosaka et al., 1993) have been reported. The estimated concentrations of PV and CB are in line with experimental data revealing the very high Ca2þ‐buffering capacities of Purkinje cells (Fierro et al., 1996; Maeda et al., 1999). The contribution of other CaBPs to the total Ca2þ‐buffering capacity of Purkinje cells is likely to be minor (Schmidt et al., 2003b). There remains one point of concern: all quantitative data (obtained using immunohistochemistry, ELISA, RIA) are based on the use of antibodies that specifically recognize the three proteins of interest. Several reports indicate that CaBP antibody recognition depends on the Ca2þ‐binding status of the investigated proteins (Winsky et al., 1996; Zimmermann et al., 2002). Hence, if the relative amounts of the Ca2þ‐bound and the Ca2þ‐free forms are different in nitrocellulose membranes (from Western blots) or in other in vitro preparations used for calibrative comparison with those in the cytoplasm of chemically‐fixed tissue sections (Kosaka et al., 1993), the ‘‘apparent’’ differences in CaBP concentrations that have been reported using different methods and by different laboratories could reflect these technical obstacles. However, in general, the agreement between reported values is fairly good.

3.2 Metal‐Binding Affinities In EF‐hand proteins, two types of Ca2þ‐binding sites are distinguished on the basis of their different selectivities and affinities for Ca2þ and Mg2þ (Celio et al., 1996). CB and CR contain only the so‐called Ca2þ‐specific sites (four functional sites in CB and five in CR; reviewed in Celio et al. (1996)). In general, the affinities of these Ca2þ‐specific sites for Ca2þ (KCa) are in the order of 10–3–10–7 M and are significantly lower for Mg2þ (KMg ¼ 10–1–10–2 M). The most accurate values for CB have been obtained in vitro by using UV‐flash photolysis of DM‐nitrophen in combination with a model‐based analysis of fluorescence transients using the Ca2þ indicator Oregon Green 488 BAPTA‐5N (Nagerl et al., 2000b). UV laser flashes of variable intensity elicit rapid, step‐like increases in [Ca2þ]i. If synthetic buffers such as EGTA or recombinant CB are added, the decay in flash‐induced fluorescence can be attributed to the presence of the Ca2þ buffer (EGTA or CB). Subsequent modeling of the transients yields the kinetic information. Using this tool, CB has been shown to possess two different types of binding sites with different KCa and kinetic Ca2þ‐ binding rates (kon): two sites are high‐affinity sites, which bind Ca2þ with a kon comparable to that of EGTA (1 107 M–1 s–1); the others are of lower affinity, with an approximately eight‐fold faster kon. However, the mathematical model did not incorporate cooperativity, a mechanism known to operate within most CaBPs, including CB (Berggard et al., 2002a). Therefore, a more refined analysis is awaited. A similar evaluation for the alternatively spliced form of CR, CR‐22k (Schwaller et al., 1995), has been undertaken, but only preliminary data are as yet available in the form of a meeting abstract (Faas et al., 2003). In a ‘‘resting’’ neuron with a [Ca2þ]i of 40–100 nM, the Ca2þ‐binding sites of CB and CR are assumed to be essentially Ca2þ‐free (less than 9% of the Ca2þ‐bound form of CB has been reported to exist in a resting cell) (Berggard et al., 2002a), and thus ready for Ca2þ binding with fast kinetics when [Ca2þ]i is raised. The second, mixed Ca2þ/Mg2þ site (two of which exist in PV) binds Ca2þ with high affinity and Mg2þ with moderate affinity in a competitive manner (dissociation constants: KCa ¼ 10–7–10–9 M; KMg ¼ 10–3–10–5 M) (see > Table 5-1). Thus, in the cytoplasm of a ‘‘resting’’ neuron, the two mixed Ca2þ/Mg2þ sites of PV are occupied principally by Mg2þ, which must dissociate before Ca2þ binding can occur. EF‐hand domains usually exist in pairs, which form a tandem domain consisting of two helix–loop– helix regions linked by a short stretch of 5–10 amino acid residues. Hence, the majority of these proteins have an even number of EF‐hand domains (six in CB and CR), the uneven number (three) for PV being rather an exception. The tandem domains are not only important for the structural stability of EF‐hand

201

BAPTA 1 (1) 160a 100–1000d,f 16–160 6–60 200g 3–9 0.04–0.4 [B] ¼ 270 mM 0.07–0.2 [B] ¼ 270 mM

EGTA 1 (1) 150a 3–10d,f 0.5–1.5 700–2000

200g 28–50

4–10 [B] ¼ 270 mM 0.7–1 [B] ¼ 270 mM

3 [B] ¼ 540 mM 0.4 [B] ¼ 540 mM

43h 16

Parvalbumin 3 (2) 150*b,c 6*b,c 0.9b 1050

5.4 [B] ¼ 540 mM 0.8 [B] ¼ 540 mM

25i 8.5

Calbindin D‐28k slow # 2 200d 10d 2 500

0.4 [B] ¼ 540 mM 0.2 [B] ¼ 540 mM

25i 1.9

Calbindin D‐28k fast # 2 500d 80d 40 25

0.007–0.07 [B] ¼ 1.35 mM 0.03–0.09 [B] ¼ 1.35 mM

25 0.3–1.1

Calretinin 6 (5) 1500**e 100–1000*** 150–1500 0.7–7

CB has six EF‐hand domains, four of which are functional: two with fast binding kinetics and two with slower binding kinetics (Nagerl et al., 2000b). In the table, the two sets of binding sites are treated as separate entities, but evidently in the intact CB molecule binding sites are in close proximity. Cooperativity of Ca2þ‐binding sites was not included in the model to calculate the different parameters. On‐ and off‐rates were derived assuming koff ¼ Kdkon *Kd and kon for PV are highly dependent on [Mg2þ]—the values stated (Schwaller et al., 2002) are estimates at physiological cytosolic [Mg2þ] (0.6–0.9 mM)

#

Ca ‐binding sites (functional) Apparent Kd (nM) (at pH 7.2) kon (mM–1 s–1) koff (s–1) ¼ Kd *kon Dwell time tdwell (ms) 1/Koff DCabuffer (mm2 s–1) Shuttle distance dshuttle (mm) √(6DCabuffer *tdwell) Capture time tcapture (ms) 1/(Kon * [B]) Capture distance dcapture (mm) √(6DCa * tcapture)

2þ

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. Table 5-1 Properties of the CaBPs, PV, CR, and CB, and of the synthetic chelators EGTA and BAPTA (modified from Dargan et al. (2004); Schwaller et al. (2002)). Published values are marked by superscript alphabets (a–j). Other values are assumptions based on experimental data and are explained below

202 Emerging functions of the ‘‘Ca2+ buffers’’ parvalbumin, calbindin D-28k and calretinin in the brain

**CR has multiple sites with different Kds—the half‐saturation value is shown ([Ca2þ]50 ¼ 1.5 mM) (Schwaller et al., 1997), but higher affinities (0.3–0.4 mM for chick CR) have been reported (Stevens et al., 1997) ***The stated kon for the estimated ‘‘fast’’ binding site(s) of CR (Edmonds et al., 2000) was used to derive the other kinetic parameters. More recent data (G. Faas, unpublished) suggests that the multiple sites of CR have differing kinetics similar to CB. The diffusion coefficients DCa buffer for PV and CB were obtained by FRAP experiments a Morris et al. (1999) b Lee et al. (2000) c Schwaller et al. (2002) d Nagerl et al. (2000b) e Schwaller et al. (1997) f Naraghi (1997) g Allbritton et al. (1992). Dwell times (tdwell), reflecting how long Ca2þ will remain bound to each buffer is defined as 1/koff. The corresponding mean distances over which the Ca2þ buffer complexes will diffuse before releasing bound Ca2þ (dshuttle) were estimated as √(6DCa buffer tdwell). The parameter dshuttle for CR was calculated assuming DCa buffer ¼ 25. Mean capture times before a Ca2þ ion binds to PV, CB, or CR were calculated tcapture ¼ 1/(konB) (Stern, 1992; Roberts, 1994), where B is the concentration of Ca2þ‐free binding sites on the buffer, assuming that Ca2þ ions in the cytosol are bound to immobile endogenous buffers for 90% of the time. Corresponding mean capture distances were estimated using the relation dcapture ¼ √6DCatcapture, assuming an apparent diffusion coefficient (DCa) of 20 mm2 s–1 for Ca2þ in the presence of immobile endogenous buffers h Schmidt et al. (2003a) i Schmidt et al. (2005) and the similar value assumed for CR is based on the similar molecular mass of CB and CR. Values for EGTA and BAPTA are based on the reported mobility (283 mm2 s–1) of another small mobile molecule, IP3


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domains but also for the binding of Ca2þ to one site that allosterically affects the affinity and the binding kinetics of the second (Nelson et al., 2002).

3.3 Metal‐Binding Kinetics Within a cell, Ca2þ‐binding kinetics (kon) can vary by more than two orders of magnitude. For proteins with Ca2þ‐specific sites implicated in very rapid biological processes, such as muscle contraction (e.g., troponin C), kon can exceed 108 M–1 s–1, whereas for the slow‐onset buffer PV, values in the order of 3 106 M–1 s–1 have been reported and are directly linked to the particular Mg2þ/Ca2þ binding sites of PV (> Table 5-1 and references cited therein). In the absence of Mg2þ, the on‐rate of Ca2þ binding to PV is very rapid (1.08 108 M–1 s–1) (Lee et al., 2000). But at the resting cytosolic [Mg2þ] levels (0.3–0.9 mM) in neurons and muscle (Li‐Smerin et al., 2001; Watanabe et al., 2001), the rate for Ca2þ binding, which is determined by the rather slow Mg2þ off‐rate (Lee et al., 2000) (> Table 5-1), is much slower. As mentioned above, kon values for CR and CB are only now emerging. But it is generally agreed that for the fastest site(s) of CR, kon probably exceed(s) 108 M–1 s–1 (Edmonds et al., 2000); the Ca2þ‐binding kinetics for CB are likely to be slower (Nagerl et al., 2000b) and those for PV the slowest of the trio. The importance of the kinetic properties of PV and CB will be discussed in > Sect. 4, where the role played by CaBPs in short‐term modulation of synaptic plasticity is addressed.

3.4 Mobility and Interaction with Ligands As a general rule, the cytosolic mobility of a molecule is proportional to its size (molecular mass). However, there are notable exceptions. In Xenopus laevis oocytes, the cytosolic diffusion rate of the intracellular second messenger inositol‐1,4,5‐trisphophate (IP3) (283 mm2 s1) (Allbritton et al., 1992) is much higher than that of Ca2þ, whose movement is impeded by the presence of slowly mobile or almost completely immobile Ca2þ buffers. The diffusion rate of Ca2þ under resting conditions (D * ¼ 13 mm2 s1) increases to 65 mm2 s1 by raising [Ca2þ]i to 1 mM, at which level the immobile buffer sites are presumably saturated. The apparent mobility of Ca2þ can also be increased by the presence of mobile Ca2þ buffers, which ‘‘shuttle’’ this cation through the sea of immobile buffers. The mobility of PV and CB in the cytoplasm of Purkinje cells has been determined by two‐photon fluorescence recovery after photobleaching (FRAP) experiments. From the recovery time constant (t) and the mean geometry of spine heads and necks of Purkinje cells, the calculated apparent diffusion coefficients (D*) are 43 mm2 s1 (Schmidt et al., 2003a) and 26 mm2 s1 (Schmidt et al., 2005) for PV and CB, respectively. As yet, no measurements have been reported for CR, but judging from the similarity of their molecular masses, values comparable to those for CB are expected (apparent Mr 28 kDa for CB and 30 kDa for CR). The manner in which a Ca2þ transient is affected by the presence of a Ca2þ buffer is also linked to the intracellular localization of the buffer, i.e., whether it is freely diffusible or bound to organelles such as the plasma membrane or the cytoskeleton. In Purkinje cells, PV behaves like a freely diffusible molecule, indicating that its main role therein is to act as a slow‐onset Ca2þ buffer modulating the spatiotemporal aspects of Ca2þ signals (Schmidt et al., 2003b). CB on the other hand, is partially immobilized within the spines and dendrites, but not within the axons of cerebellar Purkinje cells (Schmidt et al., 2005). Immobilization occurs over several seconds and is enhanced by suprathreshold synaptic activity, and can be relieved by a synthetic peptide that resembles the putative CB‐binding site of myo‐inositol monophosphatase (Berggard et al., 2002b), thereby indicating that CB binds to immobilized myo‐inositol monophosphatase. Consequently, CB has been postulated to act as an activity‐dependent sensor targeting membrane/cytoskeleton‐bound myo‐inositol monophosphatase within neurons.

3.5 Models Used to Investigate the Role of Synthetic Ca2þ Buffers and CaBPs The Xenopus oocyte model is widely used to study the function of channels, and more recently, Ca2þ signaling. The Ca2þ liberation through inositol‐1,4,5‐trisphosphate receptors (IP3R), also occurring by a


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mechanism known as Ca2þ‐induced Ca2þ release (CICR), is modulated by the mobile synthetic cytosolic Ca2þ buffers EGTA and BAPTA (Dargan et al., 2003): EGTA speeds Ca2þ signals and ‘‘disperses’’ Ca2þ waves into local ‘‘puffs,’’ whereas BAPTA slows Ca2þ responses and promotes the ‘‘globalization’’ of spatially uniform Ca2þ signals. The distinct kinetics of individual Ca2þ buffers thus influences the time course and spatial distribution of IP3‐evoked Ca2þ signals. Like EGTA, an injection or the overexpression of PV evokes Ca2þ puffs, which are inhibited by the IP3R blocker heparin. An injection of CB does not induce Ca2þ puffs, thereby indicating that CaBP‐specific kinetics is of utmost importance (John et al., 2001). In the Xenopus oocyte model, the response to CR differs in several respects from that evoked by the fast buffer BAPTA (Dargan et al., 2004). At low doses, CR induces Ca2þ puffs, which are never observed with BAPTA. This finding may reflect the different kinetic characteristics of the five Ca2þ‐binding sites of CR (Faas et al., in preparation). CaBPs not only influence the release of Ca2þ from internal stores, but also modulate the activity of Ca2þ channels, including the N‐ and P/Q‐type Ca2þ channels as they affect the processes of inactivation and facilitation, and which are intimately involved in Ca2þ influx. Modulation of the N‐type Ca2þ channels by CaM was reported first, then that of the P/Q‐type. In the former case, CaM binds to the isoleucine–glutamine motif in the carboxy terminus of the a1C subunit, and in the latter, on the a1A subunit of the channel (Lee et al., 1999; Zu¨hlke et al., 1999). In the presence of low concentrations of intracellular Ca2þ chelators, Ca2þ influx through P/Q‐type channels enhances inactivation, increases recovery from inactivation, and elicits a long‐lasting facilitation of the Ca2þ current (Lee et al., 1999). A role for PV and CB in modulating this Ca2þ‐dependent inactivation has been demonstrated in 293T cells transfected with a Cav2.1 (P/Q‐type) Ca2þ channel cDNA (Kreiner et al., 2005). Interestingly, the effects are not the same as those induced by the synthetic buffers EGTA and BAPTA. The Ca2þ‐dependent inactivation of Cav2.1 has been postulated to depend on Ca2þ microdomains located immediately beneath the plasma membrane. These are believed to be affected by the amplitude of the Ca2þ current and to be differentially modulated by PV and CB (Kreiner et al., 2005).

4

Distribution of Parvalbumin (PV), Calbindin D-28k (CB), and Calretinin (CR) in the Brain

Each of the three CaBPs has a very distinct expression pattern in the brain exemplified in the mouse temporal cortex (> Figure 5-2) (for details see figure legend). In the next section, the principal expression patterns for PV, CB, and CR are summarized.

4.1 PV The expression of PV in the CNS is restricted to the neurons (Celio et al., 1981) and by large, to a GABAergic subpopulation (Celio, 1986). The pattern of PV expression in the rat brain was first described by Celio (1990). Subsequently, many reports have appeared, dealing with its developmental patterns of expression (Hendrickson et al., 1991; Solbach et al., 1991), its expression in different species (Hof et al., 1999), and changes in its expression profile under pathological conditions (Heizmann et al., 1992) (for review see Andressen et al. (1993)). In almost all brain regions, networks of GABAergic interneurons, and in particular PV‐immunoreactive neurons, seem to be important in generating and promoting synchronous activity and are involved in producing coherent oscillations. PV‐immunoreactive neurons exert strong inhibition: In the neocortex, the chandelier and basket cells (Kawaguchi et al., 1998; Gupta et al., 2000), in the hippocampus, the axo‐axonic and basket cells (Freund et al., 1996), and in the cerebellum, the molecular layer interneurons, the stellate and basket cells (Celio, 1990; Kosaka et al., 1993), are responsible for axonic, perisomatic, and dendritic inhibition. In all these three regions consisting of many repetitive modular structures termed ‘‘microcircuits’’ (Grillner et al., 2005), the PV‐immunoreactive neurons are essential components of the networks. In thalamic structures, specific neurons express PV (Jones et al., 1989), and in particular, in the thalamic reticular nucleus (RTN) almost the entire neuronal population is PV‐immunoreactive (Seto‐Ohshima et al., 1989). In the adult rat thalamus the neurons of the reticular nucleus display PV‐immunostaining and PV‐positive fibers densely innervate most of the dorsal thalamic

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. Figure 5-2 Immunohistochemistry of mouse brain sections (temporal cortex) for the three CaBPs CB (left), PV (middle), and CR (right). The number of immunoreactive interneurons in the mouse cortex increases in the order with those of PV>CB>CR. Besides the dark‐stained population of CB‐immunoreactive interneurons, there is also a lighter‐ stained population composed of many pyramidal cells distributed throughout layers 2 and 3. Thus, the total number of cortical CB‐immunoreactive neurons is the highest of all three CaBPs

domains (Frassoni et al., 1991). Furthermore, previously unobserved PV‐immunoreactive GABAergic calyciform terminals, originating from local PV‐immunoreactive interneuronal perikarya, have been shown to synapse with large PV‐negative dendrites (Csillik et al., 2005). It has been postulated that these synapses could be instrumental in intrinsic cell‐to‐cell communications, possibly those involved in synchronizing thalamocortical oscillations (Csillik et al., 2005). Reciprocal connections between PV‐ immunoreactive interneurons are also typically encountered within microcircuits of the neocortex, the hippocampus, and the cerebellum (Grillner et al., 2005). Interestingly, these neurons are connected not only via chemical synapses, but also via electrical ones (gap junctions) involving connexin 36 (Gibson et al., 1999; Fukuda et al., 2000; Galarreta et al., 2001). In connexin‐36 knockout mice, the amplitude, but not the dominant frequency of g oscillations is reduced (Hormuzdi et al., 2001). One of the exceptions concerning the preferred relationship between GABA and PV expression is the large excitatory synapse, the calyx of Held in the medial nucleus of the trapezoid body (MNTB) of rodents. From postnatal day 6 P6 to P31, the expression of PV (and CR) increases in the presynaptic calyces, and PV is also expressed in the somata of the MNTB principal neurons (Felmy et al., 2004). The advantage of this giant excitatory synapse (the calyx of Held) in the brainstem is its accessibility to experimental manipulations: a presynaptic terminal can be patch‐clamped and the Ca2þ signals simultaneously measured (Schneggenburger et al., 2000).

4.2 CB The variety of neurons expressing CB is much greater than for PV (Celio, 1990) (for review see Andressen et al. (1993)) and there are significant differences between species (Hof et al., 1999). In rats, CB‐immunoreactivity appears on embryonic day E 14 in the CNS and the sensory organs and on E15 in the peripheral nervous system. The adult pattern of CB expression is attained before birth in most regions of the brain. In general, the pattern manifested during brain development corresponds to that in the adult organism (for further details, see Enderlin et al. (1987); Hof et al. (1999)). This situation contrasts to that for CR, which is


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often only transiently expressed during development (> Sect. 3.3). CB‐immunoreactive neurons are mainly local‐circuit neurons (interneuron type). They are most abundant in the upper cortical layers II and III. In rodents, they include the small multipolar neurons with ascending dendrites ramifying in the molecular layer, small bitufted cells, pyramid‐like cells in layer II, horizontal neurons in the molecular layer, multipolar neurons with long descending dendrites, and large double‐bouquet cells (Ferrer et al., 1992). A subpopulation of CB‐immunoreactive double‐bouquet cells has been identified in the monkey, which is also immunoreactive for CR (Zaitsev et al., 2005). CB‐immunoreactive, somatostatin‐expressing Martinotti cells have also been observed (Kawaguchi et al., 1997). For details on the coexpression of CaBPs and neuropeptides and the electrophysiological classes of interneurons, see > Figure 5-3 in Markram et al. (2004) and Toledo‐Rodriguez et al. (2005). Also, a subpopulation of pyramidal cells is weakly CB‐positive.

. Figure 5-3 Enhanced green fluorescent protein (EGFP) expression in cerebellar molecular‐layer interneurons (arrows) and Purkinje cells in PV‐EGFP mice (Meyer et al., 2002). PV‐immunoreactive basket cell terminals form an axonal plexus around the Purkinje cell main dendrite, the soma, and also make a unique complex synapse—the ‘‘pinceau’’ (arrowhead)—to the initial segment of the Purkinje cell axon (Palay et al., 1974; Korn et al., 1980). Similar to native PV expression in Purkinje cells, EGFP expression levels in transgenic mice may vary from cell to cell as seen in the three depicted Purkinje cells (PV‐EGFP mice were a kind gift from H. Monyer, Dept. of Clinical Neurobiology, Heidelberg, Germany; image courtesy of M. Chat, INSERM, UMR 8118, Paris, France)

In the hypothalamus, CB‐immunoreactive cells are widely distributed. In the hippocampus, CB‐immunoreactive interneurons are observed within all subdivisions. In the human dentate gyrus, CB is strongly expressed in granule cells, but not in CA1 pyramidal cells, unlike the situation in rodents, where CA1 cells are also CB‐immunoreactive (Sloviter et al., 1991). In the cerebellum, Purkinje cells are the only cell type expressing CB (for a review of CB, CR, and PV expression in the cerebellum, see Bastianelli (2003)).

4.3 CR In the human cortex, CR, like PV and CB, is principally expressed within a subclass of interneurons that occurs in all cortical layers, but most abundantly in layers II and III, and less frequently in layer VI. In layer I, the large horizontal (Cajal–Retzius) cells are CR‐immunoreactive (Vogt Weisenhorn et al., 1994; Belichenko et al., 1995) (for a review of CR distribution in the rat fore‐ and hindbrain, see Arai et al.

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(1991); Jacobowitz et al. (1991); Re´sibois et al. (1992)). The morphology of CR‐immunoreactive neurons ranges from ‘‘bipolar, bitufted, fusiform to double bouquet cells, whose long axes lie parallel to the radial axis of the cortex’’ (Fonseca et al., 1995). Despite the virtually identical morphological features of CR‐ immunoreactive and certain CB‐immunoreactive neurons (double bouquet cells), colocalization of CB and CR in the same neurons is rare (Schwaller et al., 1999a). Consequently, double bouquet cells are considered to be a chemically heterogeneous neuronal population (for marker studies, see Toledo‐Rodriguez et al. (2005)). In the main and the accessory olfactory bulbs, the granule, periglomerular, and mitral cells are CR‐ immunoreactive (Jacobowitz et al., 1991). CR‐positive cells are also found in the substantia nigra compacta, in the ventral tegmental area, and in the nigrostriatal and mesolimbic projections. In the hippocampus, specific CR‐immunoreactive interneurons occur within all subfields of the CA1–CA3 regions and within the dentate gyrus (Gulyas et al., 1992; Miettinen et al., 1992; Freund et al., 1996) and more than 80% of these are GABA‐ positive. GABA‐negative CR‐immunoreactive neurons are located in the hilus of the dentate gyrus and in the stratum lucidum of the CA3 subfield; they are characterized by a spiny morphology. Spine‐free CR‐immunoreactive cells are usually GABA‐positive. Hence, two morphologically and neurochemically distinct subpopulations of CR‐immunoreactive neurons exist. CR‐immunoreactive neurons of the hippocampus never coexpress PV, and rarely express CB ( Sect. 4.2. As mentioned above, the subpopulation of PV‐immunoreactive GABAergic interneurons is, in almost all brain regions, critically involved in strong perisomatic inhibition, and thus in controlling the output of the principal cells. A change in the inhibitory activity of these neurons in the neocortex has been proposed as a major mechanism underlying epileptic seizures (Mihaly et al., 1997). The role of PV in maintaining the stability of neuronal networks in vivo has been assessed in PV/ mice (Schwaller et al., 2004). Pentylenetetrazole (PTZ)‐induced seizures are more severe in PV/ than in wild‐type animals, although their onset is delayed. The latter findings accord with the observation in PV/ mice that the inhibitory effect exerted by PV interneurons of the cerebellum and hippocampus is heightened in the absence of PV. In the hippocampus, PV‐deficiency facilitates the GABAAergic current reversal induced by high‐frequency stimulation, and thus the proconvulsive GABA‐mediated depolarizing postsynaptic potential. This finding indicates that PV plays a key role in regulating the local inhibitory effects exerted by GABAergic interneurons on pyramidal neurons.

5.2 CB Knockout‐ and CB Antisense Mice Similar to PV/ mice, CB‐null mutant mice (systematic name: Calb1tm1Mpin) manifest no overt phenotype related to development, histology of the nervous system, or behavior (Airaksinen et al., 1997a). Subtle

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deficits in motor coordination implicate functional disturbances in the cerebellar pathways involving Purkinje neurons, since this is the only efferent of the cerebellar cortex, and the Purkinje cell is the only neuronal type that expresses CB in the cerebellum. To demonstrate the Purkinje‐cell specificity of the motor phenotype, Purkinje cell‐specific CB/ mice have been generated (Barski et al., 2002) by crossing a transgenic strain expressing Cre recombinase controlled by the L7/pcp‐2 gene promoter (Barski et al., 2000) with another expressing a ‘‘floxed’’ CB gene (Barski et al., 2002). Like the ‘‘global’’ CB/ mice, these Purkinje cell‐specific CB/ mice manifest an impaired motor coordination phenotype (Barski et al., 2003). In both strains, similar changes in synaptically evoked (either after parallel or climbing fiber stimulation) postsynaptic Ca2þ transients are observed: the amplitude is increased and their fast, but not their slow [Ca2þ]i decay component has larger amplitudes in CB/ mice than in wild‐type mice (Airaksinen et al., 1997a). However, the delayed metabotropic glutamate receptor‐mediated Ca2þ transients were similar to those in wild‐type animals (Barski et al., 2003). A detailed quantitative analysis of the role played by CB in the dendritic Ca2þ transients of Purkinje cells has been undertaken (Schmidt et al., 2003b). By comparing the signals from PV/ cells with those from PV/CB/ cells, the contribution of CB could be deduced and the roles of both Ca2þ buffers, as well as their concentration in Purkinje cells, could be modeled. In the absence of PV and CB, peak [Ca2þ]i amplitudes are about two‐fold higher than in wild‐ type cells, and the decay in [Ca2þ]i becomes almost monophasic. Hence, the biphasic [Ca2þ]i decay kinetics are attributable not only to the slow binding sites of PV, but also to the high‐affinity sites of CB, which are characterized by rather slow kinetics (Nagerl et al., 2000b). Interestingly, ablation of the CB gene in Purkinje cells has no effect on the LTD of Purkinje cell parallel fiber synaptic transmission, which is considered as a critical determinant of normal cerebellar function (Barski et al., 2003). This is unlike in mice deficient for components of the metabotropic glutamate receptor/IP3/Ca2þ signaling pathway, where in mice lacking type 1 IP3R the motor control impairment (severe ataxia) is associated with impairment of LTD (Matsumoto et al., 1996). In summary, motor coordination deficits in CB/ mice are probably caused by subtle disturbances in the Ca2þ signaling of Purkinje cells, which, at the network level, result in the emergence of 160 Hz oscillations (Servais et al., 2005). The role of CB in short‐term plasticity has been addressed in a newly described interneuron type, the multipolar bursting cell (Blatow et al., 2003b). On the basis of existing data, saturation of a mobile cytosolic Ca2þ buffer has been proposed as a mechanism leading to paired‐pulse facilitation (Rozov et al., 2001). More recently, saturation of CB has been shown to play a major role in paired‐pulse facilitation at CB‐ containing synapses (Blatow et al., 2003a). Washout of cytosolic constituents via the patch pipette increases the amplitude of the first response and decreases paired‐pulse facilitation, which could be restored by adding either the fast synthetic buffer BAPTA or recombinant CB. The effect is termed pseudofacilitation, since the main effect of CB (a decrease in the amplitude of the inhibitory postsynaptic potential) is on the first response in the paired‐pulse protocol. The same mechanism is also present at the CB‐immunoreactive facilitating excitatory mossy fiber CA3 pyramidal cell synapse, and experiments with CB/ mice have confirmed Ca2þ‐buffer saturation to be responsible for this presynaptic mechanism of synaptic plasticity. In many studies, a neuroprotective role for CaBPs, including CB, has been reported, details of which are summarized in > Sect. 5. In the postmortem brains of patients with mesial temporal lobe epilepsy (mTLE), which is the predominant form of the disease in adults as well as in animal models, a loss of CB from granule cells of the dentate gyrus is observed. CB’s function was addressed in both CB/ mice (Klapstein et al., 1998) and tissue obtained from patients with Ammon’s horn sclerosis (AHS) (Nagerl et al., 2000a). CB significantly increases the Ca2þ‐dependent inactivation of voltage‐dependent Ca2þ currents (ICa), thereby diminishing Ca2þ influx during repetitive neuronal firing. Addition of recombinant CB in the pipette restores ICa inactivation to levels recorded in cells with normal CB content harvested from mTLE patients without AHS. In CB/ mice, functional properties such as the altered adaptation of action potential firing and altered paired‐pulse and frequency potentiation at affected synapses are consistent with the absence of an intracellular Ca2þ‐buffer (Klapstein et al., 1998). Thus, this is a clear example, where the absence of CB contributes to increasing the resistance against excitotoxicity, and the loss of CB in dentate gyrus granule cells may be viewed as a homeostatic mechanism to protect the surviving neurons. Putative roles for CB in the visual and auditory systems have also been investigated. CB is abundant within hair cells of the inner ear and within distinct neurons of the auditory pathway, and it has been


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postulated to act as a fast buffer to speed the return of potentially toxic [Ca2þ]i levels to basal values. However, experiments with wild‐type and CB/ mice reveal no differences in the auditory brainstem‐ evoked response, in the distortion‐product otoacoustic emissions, or in noise‐induced trauma that results in hair cell loss in both strains (Airaksinen et al., 2000). These results argue against an essential role for CB either in hearing or in protecting against moderate noise‐induced inner ear traumata, at least in mice. Furthermore, the absence of CB from horizontal cells in the retina of CB/ mice has no apparent effect on the histology of this layer, which indicates that CB is not required for the structural maintenance of the differentiated retina (Wassle et al., 1998). Another approach to study the role of CB was the generation of a transgenic mouse strain producing CB antisense mRNA (Molinari et al., 1996). In these mice, impairment of plasticity (LTP in hippocampal CA1 neurons by modulation of NMDA receptors (Jouvenceau et al., 1999)) and spatial learning have been reported and linked to limiting elevations in [Ca2þ]i (Jouvenceau et al., 2002). Indeed, elevations in [Ca2þ]i effected by NMDA‐ or Kþ‐induced depolarization are prolonged in these CB‐antisense mice (Pasti et al., 1999).

5.3 CR Knockout Mice As for the other gene ablations discussed above, CR null‐mutant mice (CR/) (systematic name: Calb2tm1Map) are phenotypically only subtly compromised. The first reported alteration was impaired LTP in the hippocampus (Schurmans et al., 1997). In CR/ mice, basal synaptic transmission between the perforant pathway and granule cells and between the Schaffer commissural input and CA1 pyramidal neurons is not changed. But the induction of LTP is impaired in the dentate gyrus. The impairment is of a similar magnitude to that in CRþ/ mice indicating that a CR expression level of approximately 50% does not suffice to sustain normal LTP (Gurden et al., 1998). In the cerebellum, CR is prominently expressed in the granule cells that provide the major excitatory input to Purkinje cells via the parallel fibers, and long‐ term synaptic plasticity at the parallel fiber–Purkinje cell synapse is thought to underlie forms of motor learning also in part due to variations in [Ca2þ]i in presynaptic terminals of parallel fibers. Impairment in motor coordination tests is observed in CR/ mice and is linked to altered Ca2þ homeostasis in Purkinje cells, which is indirectly supported by the increased Ca2þ saturation of CB in these cells (Schiffmann et al., 1999). As in CB/ and PV/ mice, the firing properties of Purkinje cells are affected in alert CR/ mice. The most notable changes include an increase in the simple spike firing rate, a shortening of the complex spike duration, and a shortening of the spike pause (Schiffmann et al., 1999). These changes are not detected in brain slices, which indicates that CR is involved at the network level in cerebellar physiology. Direct patch‐clamp recordings of mature CR/ granule cells reveal these to have faster action potentials and to generate repetitive spike discharge showing an enhanced frequency increase with injected currents. Coinjection of the exogenous fast Ca2þ buffer BAPTA (150 mM) restores the wild‐type situation indicating that the absence of CR‐mediated Ca2þ buffering in CR/ cells is responsible for the observed effects (Gall et al., 2003). In alert CR/ mice, multielectrode recordings reveal the presence of 160 Hz local field potential oscillations in the cerebellar cortex (Cheron et al., 2004), similar to those observed in PV/ and CB/ mice (Servais et al., 2005). Such oscillations have never been detected in wild‐type mice. After the injection of gap‐junction‐, GABAA‐, or NMDA blockers, these 160 Hz oscillations are reversibly attenuated, which suggests that they emerge via a mechanism that synchronizes Purkinje cell assemblies (mediated by excitation of the parallel fibers) and the network of coupled molecular‐layer interneurons (stellate/basket cells). As CR‐immunoreactive granule cells are not the only CR‐expressing neurons involved in motor coordination and motor learning, the neuronal specificity of these 160 Hz oscillations has been addressed using ‘‘rescue’’ mice, where CR is selectively expressed in the granule cells (Bearzatto et al., 2006). In transgenic mice that express CR under the control of the GABAA receptor a6 promoter (Aller et al., 2003), granule cell‐specific CR expression is achieved. When these mice are crossed with CR/ mice, cerebellar granule cells remain the only neuronal subpopulation expressing CR. In alert ‘‘rescue’’ mice of this strain, granule cell excitability and Purkinje cell firing do not differ from the responses in wild‐type animals, and neither 160 Hz oscillations nor an impairment in motor coordination is observed. These results

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demonstrate that CR expression in granule cells of the cerebellar cortex is a requirement for correct computation. Therefore, fine‐tuning of granule cell excitability via Ca2þ homeostasis regulation is crucial for the coding and storage of information in the cerebellum. The same holds true for the subtle Ca2þ regulation in molecular‐layer interneurons and in CB‐expressing Purkinje cells (Servais et al., 2005).

5.4 Multitransgenic and Reporter Strains: CaMII‐PV, Thy‐1‐PV, PV‐EGFP, and Cre‐PV Ectopic CaBP expression has been reported only in two PV transgenic mouse lines. In the first mouse strain, the transgene consists of the rat CaMII promoter followed by the rat PV cDNA. PV expression is seen in the spinal cord, and additionally, the rat PV transcript is detected in liver, kidney, and muscle (Beers et al., 2001). In the second transgenic line, under the control of the Thy‐1 promoter (Chang et al., 1985), ectopic PV expression is detected within neurons of the CNS, and also in kidney, thymus, and spleen (Van Den Bosch et al., 2002). Using this model, the putative neuroprotective role of PV in striatal neurons and motoneurons has been addressed (see > Sect. 5). In one interesting reporter strain, enhanced green fluorescent protein (EGFP) is expressed in virtually all PV‐expressing neurons in various regions of the brain (Meyer et al., 2002). The protein is visible not only in the somata, but also in the dendrites and even in the axons (> Figure 5‐3). The high fidelity of expression is achieved by using an artificial bacterial chromosome (BAC) clone, which contains most (if not all) regulatory elements of the PV promoter. These mice have been used to investigate the electrical coupling between PV‐immunoreactive cells in brain slices, i.e., between dentate gyrus basket cells in the hippocampus and multipolar cells in layer II/III of the neocortex (Meyer et al., 2002). In another study, paired recordings from PV‐EGFP hippocampal basket cells were carried out to unravel the underlying mechanisms that generate g frequency oscillations (Bartos et al., 2002). The decay in unitary IPSCs at basket cell–basket cell synapses is significantly faster than that at basket cell–principal cell synapses, indicating target cell‐specific differences in IPSC kinetics. Finally, these mice have also been used for performing two‐photon targeted patching that uses two‐photon imaging to guide in vivo whole‐cell recordings of EGFP‐labeled PV‐immunoreactive interneurons in the somatosensory cortex (Margrie et al., 2003). A so‐called ‘‘knockin’’ mouse strain has also been produced, wherein an IRES‐Cre‐pA targeting cassette is integrated into the 30 ‐UTR of exon 5 of the PV gene (Hippenmeyer et al., 2005). When crossed with the appropriate floxed strain, these mice may serve to selectively express a protein in the population of PV‐immunoreactive neurons. Mouse strains with a Purkinje cell‐specific ablation of CB (Barski et al., 2003) and a selective rescue of CR in granule cells (Bearzatto et al., 2006) have been described. For the three CaBPs, CB, CR, and PV, all conceivable deletion combinations have been produced and described: CB/ CR/ (Cheron et al., 2004), CB/PV/ (Vecellio et al., 2000), and CR/PV/ (Bouilleret et al., 2000). The results obtained with these mice are described in > Sect. 5. Knockout mice for all three CaBPs have also been produced and are currently undergoing investigation (B. Schwaller et al., unpublished data). Such triple‐knockout mice are viable, fertile, and have no striking phenotype when maintained under normal housing conditions. And the histology of the brain also appears to be unaltered.

6

Alterations in CaBP Expression and Relation to Brain Pathologies

Numerous studies have reported alterations in the expression of CaBPs within diverse regions of the brain of deceased patients with specific pathologies, as well as in animal models of these diseases. These pathologies include Alzheimer’s, Huntington’s, and Parkinson’s diseases, various types of ataxia, and disorders such as epilepsy, schizophrenia, bipolar disorder, and depression (for a detailed review, see Baimbridge et al. (1992); Heizmann et al. (1992); Andressen et al. (1993); Schwaller et al. (2002)). Since the early 1990s, many more reports on this topic have been published with often contradictory results. The reason for this discrepancy is discussed next.


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6.1 Neuronal Loss vs. Loss of CaBP Immunoreactivity The overt finding may be a relatively simple one: The staining of brain sections with an antibody against CB, CR, or PV may reveal region‐specific differences between control samples and specimens derived from patients with a particular neurological disorder. The following questions arise from these observations: Does the immunostaining pattern reflect a downregulation in protein expression? Is it attributable to a loss of the neuronal population expressing this particular CaBP? If so, is there a correlation or a causal relationship between the disappearance of this neuronal population and CaBP expression? Unfortunately, many studies lack the appropriate controls to answer any of these questions with certainty. If a neuronal population expressing a particular CaBP in a given paradigm has a higher survival rate than a neighboring one, this fact is incorrectly attributed to a CaBP‐mediated neuroprotective effect. But how can one avoid making the most obvious mistakes? In general, most of the neurons expressing CB, CR, or PV selectively coexpress other neuronal markers. For example, PV‐immunoreactive neurons are, in most regions, characteristically surrounded by a special kind of extracellular matrix termed the perineuronal net (for review see Celio et al. (1994); (1998)). In PV/ mice, the number of neurons surrounded by a perineuronal net does not change, either in the cortex (Schwaller et al., 2004) or in the hippocampus (Vreugdenhil et al., 2003). Hence, the population of neurons that would express PV in wild‐type mice does not disappear in the absence of this CaBP. Other useful markers for the PV‐immunoreactive neurons are the voltage‐dependent Kþ channels Kv3.1 (Du et al., 1996) and connexin 36 (Belluardo et al., 2000). Other regional markers exist also for CB‐ and CR‐immunoreactive neurons, and those pertaining to the neocortex have been particularly well characterized (Toledo‐Rodriguez et al., 2005). The question as to whether a correlation or a causal relationship exists between the disappearance of a neuronal population and CaBP expression is experimentally demanding to answer. This issue is addressed in the next two sections.

6.2 Correlation vs. Cause vs. Secondary Adaptive Changes The putative neuroprotective roles of CB and CR in excitotoxicity have been recently reviewed (Schwaller et al., 2002), and the main conclusions are recapitulated here. In CB/ mice, the increased resistance to ischemia manifested by hippocampal neurons in vitro and in vivo (Klapstein et al., 1998) is partially attributable to an enhanced Ca2þ‐dependent inactivation of Ca2þ channels, which in turn, decreases the total Ca2þ load (Nagerl et al., 1998). This mechanism may also underlie the survival of dentate granule cells in mTLE patients with AHS (Nagerl et al., 2000a). This is in contrast to an earlier view that the absence of CB from the dentate granule cells of human epileptic patients ‘‘results in hyperexcitability of the dentate gyrus, which may then function as a motor for seizures’’ (Magloczky et al., 1997). Hence, a causal relationship between CB expression and enhanced (not decreased) vulnerability to cell death is a novel insight derived from experiments with CB/ mice. A neuroprotective role of CB was also not supported in another model based on the presence of CB‐immunoreactive neurons in the substantia nigra and the ventral tegmental areas, which were found to be more resistant to MPTP toxicity (Iacopino et al., 1992), an experimental model of Parkinson’s; the extent of neurodegeneration is identical in CB/ mice (Airaksinen et al., 1997b). The same holds true in weaver mice, a genetic model of degeneration of both midbrain dopaminergic neurons and cerebellar granule cells (Airaksinen et al., 1997b). Just as not to leave one with the impression that all studies using transgenic (knockout) mice yield consistent results, the extent of kainate‐induced excitotoxicity (a model of mTLE) was independent of the presence of the CaBPs, CB, CR, and PV (Bouilleret et al., 2000). These findings contradict others relating both to CB/ mice (Nagerl et al., 1998) and mTLE patients described above (Nagerl et al., 2000a). However, it is necessary to take into account the various changes that occur within the hippocampal GABAergic interneuron circuits, which play a central role in epileptogenesis (Magloczky et al., 2005) and which may be differentially affected in human mTLE and in murine models of the disease. Furthermore, cell survival in humans might be correlated with, but not causally related to, the absence of CB. A protective role for CB in other paradigms

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have also been reported (e.g., overexpression of CB in the striatum of rats in vivo (Yenari et al., 2001) and in vitro (D’Orlando et al., 2001)). In conclusion, the role of a particular CaBP in neuroprotection cannot be generalized; it must be carefully evaluated for each model and for each neuronal population. The data for CR are also discrepant. In vivo, CR‐containing neurons have shown to be less vulnerable than other neuronal populations to excitatory insults (Mo¨ckel et al., 1994). In several in vitro models, a neuroprotective effect of CR has been demonstrated (Pike et al., 1995; Diop et al., 1996; D’Orlando et al., 2002), although not in all (Kuznicki et al., 1996; Isaacs et al., 2000). In correlative in vivo studies, a selective sparing of CR‐containing neurons in specific regions of the brain has been demonstrated for several neurodegenerative disorders such as the striatum in Huntington’s disease (Cicchetti et al., 1996), in the substantia nigra pars compacta in Parkinson’s disease (Mouatt‐Prigent et al., 1994), and in the neocortex in Alzheimer’s disease (Sampson et al., 1997). Nevertheless, in none of these studies was a causal relationship between CR expression and increased resistance (survival) corroborated. Recent reports on the role of PV in neuroprotection using transgenic mouse models are summarized here. In CaMII‐PV mice, PV overexpression protects vulnerable motoneurons from IgG-mediated increases in [Ca2þ]i such as seen in amyotrophic lateral sclerosis (ALS) (Beers et al., 2001). Furthermore, PV expression rescues motoneurons in an animal model of familial ALS: when mice expressing mutant human Cu2þ/Zn2þ superoxide dismutase (mSOD1), an enzyme involved in free oxygen radical metabolism, are crossed with CaMII‐PV mice, there is a significant delay in the onset of the disease as compared with the mSOD1 mice (Beers et al., 2001). Thy‐PV mice ectopically expressing PV also in motoneurons were used to address whether the increase in the Ca2þ‐buffering capacity of motoneurons is protective. Previously, the selective vulnerability of motoneurons in ALS was attributed to the extremely low Ca2þ‐ buffering capacity of these cells (Palecek et al., 1999) in contrast to oculomotor neurons, which are extremely resistant in ALS models (Vanselow et al., 2000) and express high levels of PV. Cultured motoneurons in vitro from Thy‐PV mice are significantly more resistant to kainate‐induced excitotoxicity than those from control mice (Van Den Bosch et al., 2002). Furthermore, kainate‐induced Ca2þ transients mediated via Ca2þ‐permeable AMPA receptors—but not those induced by depolarization—were attenuated. The role of ectopic PV expression in motoneurons has also been investigated in vivo. Following nerve injury in neonatal rats, a large percentage of motoneurons die, probably owing to glutamate excitotoxicity, and as proposed also for motoneuron degenerative diseases such as ALS. In this study, one of the hindlimb sciatic nerves of newborn Thy‐PV‐ and wild‐type mice was crushed, and the effects on motoneuron survival were assessed 8 weeks later by retrograde labeling of the motoneurons innervating the tibialis anterior muscle (Dekkers et al., 2004). Motoneuron survival was more than two‐fold higher in the Thy‐PV mice than in the controls. However, this dramatic increase was not reflected in a significant improvement in muscle function. The putative neuroprotective role of PV in the brain has also been assessed in adult Thy‐PV mice after injecting the glutamate agonist ibotenic acid into the striatum (Maetzler et al., 2004). The result was a local loss of nerve cells and reactive astrogliosis. Contrary to the expectations of a neuroprotective role for PV, an enlarged and accelerated neurodegenerative process was observed. This finding indicates that an increase in the cytosolic Ca2þ‐buffering capacity impairs other systems involved in Ca2þhomeostasis and sequestration. The reduced mitochondrial volume in striatal neurons of Thy‐PV mice resulting from a homeostatic mechanism induced by ectopic PV expression may render these mice more vulnerable to excitotoxic stress (for details see > Sect. 5.3).

6.3 The ‘‘Ca2þ Homeostasome’’ Constitutive genetic ablation of a particular gene might induce compensatory or homeostatic mechanisms. Interestingly, in knockout mice for CB, CR, and PV, the two remaining proteins are expressed at normal levels, and in none other than the wild‐type neuronal populations. Hence, if compensatory mechanisms are at play, they are not operative at the level of the two remaining CaBPs. Indeed, there is no evidence that other CaBPs are significantly upregulated in these knockout strains. It has been postulated that either the promoter regions of other CaBPs are permanently inactivated in the neuronal population lacking a


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particular CaBP or that the relevant parameters for the two remaining CaBPs (affinities, kinetics, and diffusion) are not suited to compensate for the missing one (Schwaller et al., 2002). Nevertheless, putative adaptive changes have been reported. The spine morphology of Purkinje cells is selectively affected by the absence of CB (increased length and volume), but not by that of PV (Vecellio et al., 2000). On the other hand, the volume of mitochondria, organelles involved in Ca2þ sequestration and acting as a transient Ca2þ store, is almost twice as large in fast‐twitch muscles of PV/ mice (Chen et al., 2001). The reverse effect is observed in neurons that express PV ectopically (Maetzler et al., 2004). Within striatal neurons, which do not normally express PV, the mitochondrial volume is reduced by almost 50%, which accounts for the heightened excitotoxic injury provoked by a local injection of ibotenic acid into Thy‐PV mice. In several neurological diseases, including schizophrenia, a decrease in the population of prefrontal GABAergic PV‐ immunoreactive neurons has been reported (Lewis et al., 2005). In the same subpopulation, mRNAs for the 67‐kDa isoform of glutamate decarboxylase (GAD67), an enzyme for GABA synthesis and for the GABA membrane transporter (GAT1), were decreased, which resulted in impaired neurotransmission. Deficient neurotrophin signaling via the tyrosine kinase receptor B was postulated as the underlying pathogenetic mechanism. The downregulation of PV observed under these conditions is viewed as a compensatory mechanism to increase GABAergic transmission (Lewis et al., 2005). This postulate is moreover based on the finding that repetitive inhibitory postsynaptic currents are facilitated, and the power of g oscillations is increased in the hippocampus of PV/ mice (Vreugdenhil et al., 2003), indicating increased inhibition. Hence, the ‘‘Ca2þ buffers,’’ CR, CB, and PV, constitute an integral part of the precisely tuned system involved in Ca2þ homeostasis and Ca2þ signaling. Elimination of a CaBP does not induce the obvious compensatory mechanism (i.e., the upregulation of another Ca2þ buffer), but leads to more subtle changes at the level of cell morphology or within specific Ca2þ‐uptake or ‐release systems. Apparently, CaBP‐ deficient cells make use of the components of the ‘‘Ca2þ‐signaling toolkit’’ (Berridge et al., 2003), which they adapt to conform to the situation prevailing in ‘‘normal’’ wild‐type cells.

7

Conclusion

The three EF‐hand Ca2þ‐binding proteins, CB, CR, and PV, which, in classical terminology, are referred to as ‘‘Ca2þ buffers,’’ are integral components of Ca2þ‐signaling and Ca2þ homeostatic mechanisms in specific CNS neurons. The properties of PV, particularly its great mobility and inability to interact with other cellular components, render this molecule the prototype of a slow‐onset Ca2þ buffer. While most of the functions of CB and CR can be explained in terms of the fast Ca2þ‐buffering kinetics of their Ca2þ‐specific binding sites, their additional roles as Ca2þ modulators are now starting to emerge. The best‐characterized interaction is the binding of CB to the enzyme myo‐inositol monophosphatase, which links IP3‐ and Ca2þ‐ signaling pathways, at least in Purkinje cells. Novel insights into the important functions of CB, CR, and PV in the CNS have been obtained from genetically modified mice using both in vitro and in vivo approaches. Further progress in the field is expected using even more subtle genetic tools, such as subpopulation‐specific inducible CaBP ‘‘knockout’’ and ‘‘knockin’’ strains, as well as multitransgenic mice with alterations to several components of the neuronal Ca2þ‐signaling system. These models should help us tackle the tremendous complex mechanisms underlying intraneuronal Ca2þ signaling and Ca2þ homeostasis.

Acknowledgments The help of M. Celio, Fribourg, Switzerland and I. Llano, Paris, France, for the critical reading of the manuscript is highly appreciated. > Figure 5-3 was provided by M. Chat, INSERM, UMR 8118, Paris, France. This work was partially supported by a grant from the Swiss National Science Foundation (no. 3100A0‐100400/1 to B. S.).

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Actin, Actin‐binding Proteins and Myosins in Nervous System

R. Ishikawa

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225

2 2.1 2.2 2.2.1 2.2.2 2.3 2.3.1 2.3.2

Actin in Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Actin Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Actin Isoforms in Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Differential Function of Actin Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Differential Expression of Actin Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226 Polymerization of Actin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226 ‘‘Nucleation,’’ ‘‘Elongation,’’ and ‘‘Equilibration’’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226 Polarity of F‐Actin and Treadmilling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226

3 3.1 3.1.1 3.1.2 3.2 3.2.1 3.2.2 3.3 3.3.1 3.3.2 3.3.3 3.4 3.4.1 3.4.2 3.5 3.5.1 3.5.2 3.5.3

Actin‐Binding Proteins in Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228 Actin‐Severing/Capping Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228 Gelsolin Superfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228 ADF/Cofilin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 G‐Actin Binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 Profilin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 Thymosin b . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 230 Actin Polymerization‐Promoting Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 Arp2/3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 N‐WASP and WAVE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 mDia1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 Actin‐Crosslinking Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 Fascin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 a‐Actinin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 Side‐Binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 Tropomyosin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 Caldesmon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234 Drebrin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234

4 4.1 4.2 4.2.1 4.2.2 4.2.3 4.3 4.4 4.5 4.5.1 4.5.2

Myosin Superfamily in Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 Common Properties of Myosin Superfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 Conventional Myosin (Myosin II) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 Differential Expression of Myosin II Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 Differential Function of Myosin II Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 Subcellular Localization of Myosin II Isoforms in Growth Cone . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236 Myosin I . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236 Myosin V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236 Other Minor Myosins in Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 Myosin III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 Myosin VI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237

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4.5.3 Myosin VII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 4.5.4 Myosin IX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 4.5.5 Myosin X . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 5

Conclusion: Possible Roles of Actin, Actin‐Binding proteins, and Myosins in Living Cells . . 238


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Abstract: In this chapter, I summarize the basic properties of actin, including polymerization process, polarity of F-actin, and treadmiling, which are common in nerve cells as well as other nonmuscle and muscle cells. The properties are modulated by varieties of actin-binding proteins. Thus, I pick up some major actin-binding proteins expressing in nerve cells, and summarize (i) how these proteins affect actin properties, and (ii) possible functions of these proteins in nervous system. Finally, both conventional and unconventional myosins expressing in nerve cells are also shown, and discussed in the possible roles in actin dynamics in nerve cells. List of Abbreviations: G-actin, globular actin; F-actin, filamentous actin; ATP, adenosine 5´-triphosphate; ADP, adenosine 5´-diphosphate; Cc, critical concentration; NMDA, N-methyl-D-aspartate; DRG, dorsal root ganglion; ADF, actin depolymerizing factor; Arp, actin related protein; LPA, lysophosphatidic acid; GTP, guanosine 5´-triphosphate

1

Introduction

Actin is one of the most abundant proteins in the nervous system and forms a framework in neuronal cells. Actin polymerizes head‐to‐tail and forms long filaments. Sometimes the filaments are cross‐linked, forming a variety of structures such as flat‐sheet networks, tight bundles, and membrane‐bound networks, which determine and support cell shape. A second important function of actin is to supply rails for myosins. Myosins are the motor proteins that translocate along actin filaments, in a process powered by energy from hydrolysis of ATP. The mechanical force thus generated is used in many neuronal phenomena, including growth cone motility, activity‐dependent motility of dendritic spines, and vesicle transport along axons and dendrites. From the 1980s onward, a variety of actin‐binding proteins have been discovered, and evidence suggests that they regulate actin organization and actin–myosin interaction. During the same period, new types of myosin molecules have also been discovered. In this chapter, I focus on the molecular properties of actin, myosin, and actin‐binding proteins. Cross talk between these proteins and possible roles of these proteins in nerve cell dynamics are also discussed.

2

Actin in Nervous System

2.1 Actin Structure Actin is a globular molecule composed of 375–377 amino acids. Monomer actin, also known as globular actin (G‐actin), has two structurally similar domains (the result of gene duplication) and divalent cation‐ binding and ATP (or ADP)‐binding pockets in the hinge region of the two domains (Kabsch et al., 1990). At physiological salt concentrations, they polymerize head‐to‐tail and form a double‐helical complex (F‐actin) with a diameter of 5–9 nm (Holmes et al., 1990). The half pitch of the helix is 37 nm, and contains 13 actin molecules.

2.2 Actin Isoforms in Nervous System 2.2.1 Differential Function of Actin Isoforms Higher organisms have six major isoforms of actin; a‐skeletal muscle type, a‐cardiac muscle type, a‐(vascular) smooth muscle type, g‐smooth muscle type, b‐nonmuscle type, and g‐nonmuscle type. All mammalian and avian species have completely identical amino acid sequences of isoforms, indicating that actin is one of the most conserved proteins. Several reports suggest that there are functional differences

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between actin isoforms. Affinities of profilin (Larsson and Lindberg, 1988; Ohshima et al., 1989) and thymosin b (Weber et al., 1992), which are G‐actin‐binding proteins, for nonmuscle actin (mixture of b‐nonmuscle actin and g‐nonmuscle actin) are stronger than their affinities for a‐skeletal muscle actin. When cDNA of actin isoforms is transfected into cultured cardiomyocytes (von Arx et al., 1995) and cultured smooth muscle cells (Mounier et al., 1997), muscle‐type actin is incorporated into sarcomeres and nonmuscle actin localizes in the periphery. Furthermore, transfection of cDNA of nonmuscle actins markedly affects the shape of muscle and nonmuscle cells, whereas transfection of cDNA of muscle actins has no such effects (von Arx et al., 1995; Mounier et al., 1997). These previous findings suggest that there are functional differences among actin isoforms and further biochemical characterization is needed to clarify this issue.

2.2.2 Differential Expression of Actin Isoforms b‐Nonmuscle actin and g‐nonmuscle actin are expressed in nerve cells as well as in other nonmuscle cells. In rat brain, expression of b‐nonmuscle actin is high in E18 embryonic stages, and gradually decreases to half‐maximum level in the adult stage (Weinberger et al., 1996). b‐Nonmuscle actin localizes in the axons, growth cones and soma of developing neurons, but is absent from axons of mature neurons (Rochlin et al., 1995; Weinberger et al., 1996). In contrast, g‐nonmuscle actin is expressed at a constant level throughout development, and evenly distributed throughout the cell bodies of neurons (Weinberger et al., 1996).

2.3 Polymerization of Actin 2.3.1 ‘‘Nucleation,’’ ‘‘Elongation,’’ and ‘‘Equilibration’’ When a sufficient amount of G‐actin is introduced at physiological salt concentrations, most of the molecules polymerize and form F‐actin. Polymerization starts with an initial lag phase (so‐called nucleation phase), followed by a linear growth phase (so‐called elongation phase), and concludes with a final ‘‘equilibrium’’ phase (> Figure 6-1a). Polymerization is initiated by spontaneous collision between two G‐actin molecules (> Figure 6-1b). A dimer actin complex is formed after collision, which is not stable and can be easily separated. Only when monomer G‐actin collides with dimer actin to form a trimer does the actin complex become stable. Thus, double collisions between actin molecules in a short period are necessary to initiate actin polymerization. The probability of double collisions is much lower than that of single collisions; this is manifested as the initial lag phase of polymerization. Once the trimer is formed, the next step in the polymerization process (the ‘‘elongation phase’’) begins (> Figure 6-1b). When G‐actin binds to one end of F‐actin, the G‐actin remains stably attached at that end. The rate constant of this reaction is independent of the length of F‐actin; i.e., the elongation speeds of short F‐actin and long F‐actin are identical with an equal concentration of G‐actin. However, the elongation rate differs quite markedly between the barbed end and pointed end; i.e., the barbed end grows much faster than the pointed end. In the final step of polymerization, the free G‐actin concentration gradually decreases, and at a certain G‐actin concentration there is an ‘‘equilibration’’ between addition of G‐actin to the ends of F‐actin and loss of G‐actin from the ends of F‐actin (> Figure 6-1b). This G‐actin concentration is called the ‘‘critical concentration’’ (Cc) of actin polymerization. After ‘‘equilibration,’’ the final concentration of F‐actin depends on the total concentration of actin, but the amount of G‐actin remains constant (i.e., it is equal to Cc).

2.3.2 Polarity of F‐Actin and Treadmilling However, even in this ‘‘equilibrium’’ state, there is no equilibration between association and dissociation at the ends of F‐actin in the presence of ATP. Actin molecules can bind to both ATP and ADP, and the Cc of ATP‐bound G‐actin (ATP/G‐actin) is much lower than the Cc of ADP‐bound G‐actin (ADP/G‐actin).


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. Figure 6-1 Polymerizaton of actin. (a) polymerization profile. In the presence of physiological salt concentration, actin starts with an initial lag phase ((1) nucleation phase), followed by a linear growth phase ((2) elongation phase), and concludes with (3) a final ‘‘equilibrium’’ phase. (b) Schematic illustration of each phase

Furthermore, when ATP‐bound actin is incorporated into F‐actin, the ATP is hydrolyzed to ADP. In the ‘‘equilibrium’’ phase, the ATP hydrolysis rate of incorporated actin is lower than the rate of association of ATP/G‐actin into the barbed end, but greater than the rate of association of ATP/G‐actin into the pointed end. This results in the polar distribution of actin molecules in F‐actin, with ATP/actin predominant at the

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barbed end and ADP/actin predominant at the pointed end. This polar distribution reflects the difference in Cc between the pointed end (Cc(p); 1 mM) and barbed end (Cc(b); 0.1 mM). Because Cc(b) < [G‐actin] (¼Cc) < Cc(p) in the ‘‘equilibrium’’ state, the barbed end is always polymerizing and the pointed end is always depolymerizing. The rate of incorporation of G‐actin into the barbed end of F‐actin is equal to the rate of dissociation of G‐actin from the pointed end of F‐actin, so that the length of F‐actin is always constant. However, the position of each incorporated actin molecule gradually moves from the barbed end to the pointed end, in a process known as ‘‘treadmilling.’’

3

Actin‐Binding Proteins in Nervous System

A variety of actin structures are present in nerve cells, including flat networks of F‐actin lamellipodia, tight bundles of F‐actin filopodia, and supporting networks of cell cortex. All of these structures contain not only actin molecules but also a wide variety of actin‐binding proteins.

3.1 Actin‐Severing/Capping Proteins 3.1.1 Gelsolin Superfamily Gelsolin, an 82‐kDa globular protein first identified in rabbit macrophages (Yin and Stossel, 1979), is expressed in the brain and concentrated in oligodendrocytes and Schwann cells (Tanaka and Sobue, 1994). Gelsolin affects the organization of actin via the following (> Figure 6-2) (for review; Sun et al., 1999): (1) F‐actin severing activity (> Figure 6-2a), in which gelsolin binds to the middle of F‐actin and severs the filament in the presence of Ca2þ ions; (2) F‐actin capping activity (> Figure 6-2b), in which, after severing the filament, gelsolin remains at the barbed end of the severed‐filament and inhibits polymerization at that end; this capping activity is abolished by PIP2 (Janmey and Stossel, 1987); and (3) nucleation‐promoting activity (> Figure 6-2c), in which gelsolin binds two G‐actin molecules in the presence of Ca2þ. Like the trimer complex of actin molecules, the gelsolin/actin complex acts as a ‘‘seed’’ for elongation. Because affinities of gelsolin for G‐actins are much higher than the affinity of G‐actin for G‐actin, gelsolin accelerates the nucleation process of actin polymerization. Consequently, a large number of short F‐actin filaments are formed (> Figure 6-2). Primary culture of hippocampal neurons from gelsolin‐knockout mice exhibits an increased number of filopodia with a slow filopodia retraction speed, suggesting that gelsolin affects the growth cone dynamics (Lu et al., 1997). Evidence also suggests that gelsolin affects Ca2þ influx and apoptosis of neurons. Ca2þ influx into cultured hippocampal neurons after NMDA treatment is greatly enhanced in gelsolin‐ knockout mice, and the survival rate after treatment with glutamate is lower in gelsolin‐knockout mice (Furukawa et al., 1997). Some drug‐induced apoptosis of cerebral cortex neurons is greatly enhanced in gelsolin‐knockout mice, and this enhancement is abolished by actin‐depolymerizing drugs (Harms et al., 2004). Adseverin, an 81‐kDa gelsolin‐like protein that has actin‐severing, ‐capping, and ‐nucleating activities (Maekawa et al., 1989) and shares six homologous domains with gelsolin (Nakamura et al., 1994), is expressed in neural tissues (Sakurai et al., 1990). However, its function in neural issues is still unclear. Advillin, a 92‐kDa gelsolin‐like protein, shares six homologous domains with gelsolin, and has a carboxy‐ terminal headpiece (Marks et al., 1998). Advillin is expressed in the brain and is especially concentrated in dorsal root ganglia (DRG) and cerebral cortex (Marks et al., 1998). Advillin associates with the scavenger receptor SREC‐I in DRG neurons, and coexpression of advillin and SREC‐I causes formation of process‐like structures in a neuronal cell line (Shibata et al., 2004), but the molecular mechanism of formation of these structures is still unclear. CapG, a 41‐kDa protein that has actin‐capping and ‐nucleating activities but lacks actin‐severing activity, shares three N‐terminal homologous domains with gelsolin (Dabiri et al., 1992). It is expressed in the developing brain cortex (Arai and Kwiatkowski, 1999), but its function in neural development is unclear.


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. Figure 6-2 (a) Severing, (b) capping, and (c) nucleation promoting activity of gelsolin. Note that all the activities cause the formation of a large number of short F‐actin filaments (see text)

3.1.2 ADF/Cofilin Actin‐depolymerizing factor (ADF) and cofilin are 15–22‐kDa proteins that sever F‐actin in a pH‐dependent manner and form a protein family because of the similarity between their amino acid sequences (for review; Moon and Drubin, 1995; Maciver and Hussey, 2002). Although their amino acid sequence is not homologous with gelsolin, the tertiary structure of ADF/cofilin is quite similar to that of one of the six repetitive domains of gelsolin (Hatanaka et al., 1996). ADF/cofilin binds both F‐actin and G‐actin at a ratio of 1:1. ADF/cofilin binds much more strongly to ADP‐actin than to ATP‐actin and increases the rate constant of dissociation of actin from the pointed end, resulting in acceleration of the ‘‘treadmilling’’ speed by 25‐fold (Carlier et al., 1997). Phosphorylation of ser‐3 of ADF/cofilin by LIM kinase inactivates its actin‐binding activity (Arber et al., 1998; Yang et al., 1998). ADF/cofilin is strongly expressed in the brain (Moriyama et al., 1990). ADF/cofilin is concentrated in actin‐rich structures such as lamellipodia and filopodia in the growth cone (Bamburg and Bray, 1987) suggesting that it affects actin dynamics. Indeed, inhibition of phosphorylation of ADF/cofilin suppresses semaphorin‐3A‐induced growth cone collapse (Aizawa et al., 2001).

3.2 G‐Actin Binding Proteins 3.2.1 Profilin Profilin is a 12–19‐kDa G‐actin‐binding protein purified from a variety of tissues including brain tissue (Blikstad et al., 1980, Nishida et al., 1984). It binds to G‐actin with a stoichiometry of 1:1, and accelerates the

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exchange reaction from ADP‐bound G‐actin to ATP‐bound G‐actin (Goldschmidt‐Clermont et al., 1992), resulting in the promotion of polymerization at the barbed end of F‐actin (Pantaloni and Carlier, 1993). It should be noted that this effect occurs only when the barbed ends of F‐actin are free (> Figure 6-3a). In the presence of barbed‐end capping proteins such as gelsolin, profilin acts as a sequester protein (> Figure 6-3b) i.e., profilin depletes the free form of G‐actin in solution, resulting in F‐actin depolymerization that maintains the amount (Cc) of G‐actin (Carlsson et al., 1977). Profilin binds to PIP2, and thus loses its actin‐binding activity (Lassing and Lindberg, 1985). Furthermore, as discussed below, profilin binds to a variety of proline‐rich proteins including N‐WASP (Suetsugu et al., 1998) and mDia (Watanabe et al., 1997). Evidence suggests that in association with these proteins profilin enhances actin dynamics in neuronal cells.

. Figure 6-3 Profilin promotes actin polymerization (a) in the absence of gelsolin, whereas it causes actin depolymerization (b) in the presence of gelsolin. ‘‘ATP‐actin’’ and ‘‘ADP‐actin’’ indicate the actin molecules associated with ATP and ADP, respectively, in their nucleotide‐binding pocket

3.2.2 Thymosin b Thymosin b is a 5‐kDa protein that is widely distributed in vertebrates, including the developing nerve system (Lugo et al., 1991). It binds to G‐actin at a ratio of 1:1 and causes F‐actin depolymerization, which maintains the amount of G‐actin in solution, because the actin/thymosin b complex does not polymerize into F‐actin (Safer et al., 1990). Thymosin b inhibits the dissociation of adenine nucleotide from G‐actin, and thus inhibits the ADP/ATP exchange of depolymerized G‐actin (Goldschmidt‐Clermont et al., 1992). Thymosin b is


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thought to be a main factor in the maintenance of G‐actin pools in cells. Knockout of thymosin b expression by antisense RNA causes disruption of brain development in zebra fish (Roth et al., 1999).

3.3 Actin Polymerization‐Promoting Proteins 3.3.1 Arp2/3 Arp2/3 complex, a 220‐kDa macromolecular assembly, is composed of two actin‐related proteins (Arp2 and Arp3) whose amino acid sequences are similar to that of actin, and consists of five different subunits with molecular masses of 40 kDa, 34 kDa, 20 kDa, 21 kDa, and 16 kDa (for reviews; Weaver et al., 2003; Millard et al., 2004). It localizes at the leading edge of migrating cells including neuronal growth cones, and is thought to play a key role in the formation of branching actin networks in lamellipodia (Svitkina and Borisy, 1999) (> Figure 6-4). The activated form of Arp2/3 complex can bind to the side of F‐actin and provides a new nucleation site for actin polymerization (Volkmann et al., 2001).

3.3.2 N‐WASP and WAVE N‐WASP is a 65‐kDa neural protein that is homologous to the Wiskott–Aldrich syndrome protein (Miki et al., 1996), greatly promoting actin polymerization in association with the Arp2/3 complex (for reviews; Takenawa and Miki, 2001; Millard et al., 2004). N‐WASP contains one Arp2/3 complex‐binding site, two G‐actin‐binding sites, one profilin‐binding site, one CDC42‐binding site, and one PIP2‐binding site. In the inactivated form of N‐WASP, the Arp2/3 complex‐binding site and actin‐binding sites are covered by its internal inhibitory domain. When CDC42 and PIP2 bind to N‐WASP, both sites are uncovered and N‐WASP becomes active (Kim et al., 2000; Rohatgi et al., 2000). WAVE (also called Scar) is a 54–62‐kDa protein whose C‐terminal half domain is homologous to that of N‐WASP (Machesky and Insall, 1998; Miki et al., 1998). It localizes at the leading edge of migrating growth cones. WAVE contains one Arp2/3 complex‐binding site, one G‐actin‐binding site, and one profilin‐binding site, but lacks a CDC42‐binding site, PIP2‐binding site, and internal inhibitory domain. WAVE promotes actin polymerization in a manner similar to that of N‐WASP, but the regulatory system is different (Takenawa and Miki, 2001; Millard et al., 2004). Evidence suggests that activator of WAVE (IRSp53; Miki et al., 2000) and inhibitory complex of WAVE (PIR121þNap125þAbi2; Eden et al., 2002) regulate WAVE activity via rac‐1.

3.3.3 mDia1 The mammalian homolog of Drosophila diaphanous 1 (mDia1) is a 139‐kDa actin‐, profilin‐, and Rho‐ binding protein that shares two homologous domains with formin (Watanabe et al., 1997). It binds to the barbed end of F‐actin and promotes actin polymerization, which is enhanced by profilin (Moseley et al., 2004). Unlike Arp2/3, evidence suggests that mDir1 moves processively along the barbed end during polymerization in living cells (Higashida et al., 2004). RNAi knockdown of mDir1 inhibits axon elongation in cultured cerebellar granule neurons (Arakawa et al., 2003).

3.4 Actin‐Crosslinking Proteins 3.4.1 Fascin Fascin is a 53–57‐kDa actin‐bundling protein expressed in a variety of tissues including brain tissue. At the cellular level, it localizes in dynamic actin structures such as filopodia and microspikes (for review; Edwards

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. Figure 6-4 Formation of branching F‐actin network by Arp2/3 and N‐WASP Complex of Arp2/3 and N‐WASP binds to the barbed end or middle of F‐actin, and provides a new nucleating site for actin polymerization. Arp2/3 remains with F‐actin after polymerization, but N‐WASP dissociates from F‐actin. Dissociated N‐WASP binds to the free form of Arp2/3, and the complex binds to the F‐actin again

and Bryan, 1995; Kureishy et al., 2002). Fascin binds to F‐actin with a stoichiometry of 1 fascin molecule to 3 to 5 actin molecules, and cross‐links F‐actin to form tight F‐actin bundles (Edwards and Bryan, 1995). Reconstituted actin bundles formed by fascin and bundles observed in filopodia in growth cones have quite similar features. The distance between cross‐linked filaments in the bundle range from 8 to 9 nm, and all the filaments in the bundle point in the same direction (Ishikawa et al., 2003) (> Figure 6-5a). Furthermore, knockdown of fascin expression by antisense RNA causes loss of filopodia (Edwards and Bryan, 1995) suggesting that fascin is a bundling factor of F‐actin in filopodia of growth cones. Fascin loses its actin‐ binding activity when C‐kinase phosphorylates the ser‐39 of fascin (Ono et al., 1997) or when F‐actin filaments are covered by tropomyosin and caldesmon (Ishikawa et al., 1998).


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. Figure 6-5 Actin bundles formed by (a) fascin and (b) by a‐actinin

3.4.2 a‐Actinin a‐Actinin is a rod‐shaped, homodimeric actin‐cross‐linking protein with a subunit molecular mass of 94–103 kDa. The subunit is composed of three domains: an actin‐binding domain in its N‐terminal; a spectrin‐like rod domain; and two EF‐hand structures in its C‐terminal. The subunits associate antiparallel via their rod domains, and the dimer thus has two actin‐binding sites at each end. The distance between cross‐linked F‐actin filaments is 36 nm (Meyer and Aebi, 1990) (> Figure 6-5b). In mammals, a‐actinin is classified into four groups according to the amino acid sequence: a‐actinin‐1 to ‐4. a‐actinin‐1, a‐actinin‐2, and a‐actinin‐4 are expressed in different patterns in nerve cells. a‐actinin‐1 localizes mainly in adhesion plaques, adherence junctions, and at the end of stress fibers (Lazarides and Burridge, 1975), whereas a‐actinin‐4 localizes in stress fibers and the nucleus (Honda et al., 1998). a‐actinin‐1 binds to integrin b1 (Otey et al., 1990) and a‐catenin (Knudsen et al., 1995), and evidence suggests that it cross‐links F‐actin via adherence proteins. The actin‐binding activity of a‐actinin‐1 is inhibited by the binding of Ca2þ to the EF‐hand (Burridge and Feramisco, 1981). a‐actinin‐2, which is mainly expressed in muscle, binds to the NMDA receptor in vitro, and colocalizes with NR1 in the dendritic spines of hippocampal cultured neurons (Wyszynski et al., 1997). a‐actinin‐2 also binds to the adenosine A2A receptor (Burgueno et al., 2003). a‐actinin‐2 may serve as an anchor for these receptors in their association with F‐actin.

3.5 Side‐Binding Proteins 3.5.1 Tropomyosin Tropomyosin is a rod‐shaped, homodimeric actin‐binding protein with subunit mass of 29–33 kDa (for reviews; Pittenger et al., 1994, Gunning et al., 1998). In mammals, tropomyosins are produced by a multigene family, and about 20 isoforms are produced by alternative splicing (Pittenger et al., 1994). They are classified into two types: a higher‐molecular‐weight type (about 280 amino acids) that contains seven actin‐interacting sites and has high affinity for F‐actin; and a lower‐molecular/weight type (about 240 amino acids) that contains six actin‐interacting sites, and thus has low affinity for F‐actin (Matsumura and

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Yamashiro‐Matsumura, 1985). The tropomyosin isoforms TMBr3, TM4, and TM5, all of which belong to the lower‐molecular‐weight type, are mainly expressed in the brain, and at least seven more isoforms are also expressed (Gunning et al., 1998). Tropomyosin binds to the grooves of F‐actin filaments at a stoichiometry of 1:7, and stabilizes F‐actin filaments. Also, it inhibits the F‐actin‐capping and ‐severing activities of gelsolin (Ishikawa et al., 1989), the actin‐depolymerizing activity of ADF/cofilin (Bernstein and Bamburg, 1982), and the actin‐bundling activity of fascin (Ishikawa et al., 1998) (> Figure 6-6). . Figure 6-6 Tropomyosin affects the binding of other actin‐binding proteins. X and O indicate the inhibition and activation, respectively, of the binding to F‐actin

3.5.2 Caldesmon Caldesmon is a rod‐shaped actin‐, calmodulin‐, and tropomyosin‐binding protein that has a molecular mass of 87–89 kDa in smooth muscle cells and 60 kDa in nonmuscle cells including neurons (for review; Sobue and Sellers, 1991; Matsumura and Yamashiro, 1993). Caldesmon loses its actin‐binding activity when Ca2þ/ calmodulin binds to caldesmon (Sobue et al., 1981) and when caldesmon is phosphorylated by CDC2 kinase (Yamashiro et al., 1991). Caldesmon increases the affinity of tropomyosin for F‐actin and enhances protection of tropomyosin from gelsolin (Ishikawa et al., 1989). It also inhibits Arp2/3‐mediated actin nucleation (Yamakita et al., 2003) and actin‐activated ATPase activity of myosin II (Sobue and Sellers, 1991).

3.5.3 Drebrin Drebrin is a rod‐shaped actin‐binding protein that is mainly expressed in the brain (for review; Shirao, 1995). Two drebrin isoforms are produced by alternative splicing: an embryonic type (drebrin E) that is present in large amounts throughout the cell bodies of developing neurons; and an adult type (drebrin A) that contains 45 extra amino acids in the middle of the molecule and is concentrated in dendritic spines (Hayashi et al., 1996). Overexpression of drebrin causes changes in the shape of dendritic spines in cultured cortical neurons (Hayashi and Shirao, 1999) suggesting that drebrin is a key molecule in the morphology of dendritic spines. Drebrin binds to F‐actin with a stoichiometry of 1:5 and inhibits the actin‐binding activity of tropomyosin, a‐actinin, and fascin (Ishikawa et al., 1994; Sasaki et al., 1996). Drebrin also binds to


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profilin (Mammoto et al., 1998), although its effects on profilin activities are unclear. Drebrin inhibits the actin‐activated ATPase activity of myosin II and in vitro sliding of actin along myosin II (Hayashi et al., 1996).

4

Myosin Superfamily in Nervous System

4.1 Common Properties of Myosin Superfamily Myosins (myosin superfamily) are a group of motor proteins that generate mechanical force by hydrolyzing ATP and interact with actin filaments. The power thus obtained is used in a variety of cellular movements, including muscle contraction, cell migration, cytokinesis, vesicle transport, growth cone motility, and morphological changes of dendritic spines. Myosins are classified into 18 types, class I through XVIII, according to the protein sequence (Sellers, 2000; Berg et al., 2001). Typical myosin is composed of one or two heavy chains (HCs) and one to six pairs of light chains (LCs). HC has at least three domains: a ‘‘head domain’’ that contains an actin‐binding site and an ATP‐binding site, and which has motor activity; a ‘‘neck domain’’ that contains 1– 6 repeats of LC‐binding sites called IQ motifs (which contain the amino acid sequence IQXXXRGXXXRK), and which regulates the motor activity coupled with LC; and a ‘‘tail domain’’ that is responsible for self‐assembly and association with vesicles and cytoskeletal components. The amino acid sequences of the head domains are highly conserved throughout the myosin superfamily, whereas the sequences of the tail domains are quite diverse. The coiled‐coil structure is sometimes found in the tail region of HC or between the neck and tail. It has been hypothesized that such myosins are double‐headed myosins, because the HC can form a dimer in the coiled‐coil structure. However, only myosin II and myosin V have been confirmed to be double‐headed by electron microscopy. Major myosins found in nerve cells are myosin I, myosin II, and myosin V. Myosin III, myosin VI, myosin VII, myosin IX, myosin X, and myosin XV are also expressed in nerve cells (for review; Brown and Bridgman, 2004).

4.2 Conventional Myosin (Myosin II) 4.2.1 Differential Expression of Myosin II Isoforms Myosin II is the best‐known muscle‐type myosin, which was called ‘‘myosin’’ until the unconventional myosins were discovered. Myosin II is composed of one pair of HCs (200–240 kDa), two distinct pairs of LCs (14–20 kDa), regulatory light chains (RLCs), and essential light chains (ELCs). In neuronal cells and other nonmuscle cells, two HC genes of myosin II, myosin IIA and myosin IIB, are expressed. Myosin IIB is dominantly expressed in neuronal cells. Approximately 80% of myosin II in the cerebrum, cerebellum, brain stem, and spinal cord is the isoform myosin IIB (Takahashi et al., 1992). Myosin IIB can have a 10‐amino‐acid insertion near the ATP‐binding site of the HC (myosin IIB1) or a 21‐amino‐acid insertion near the actin‐binding site of the HC (myosin IIB2); myosin IIB1 and myosin IIB2 are produced by alternative splicing in the vertebrate brain (Takahashi et al., 1992). The levels of expression of types of myosin IIB in the human cerebrum are as follows: noninsertion type> myosin IIB1> myosin IIB2. In contrast, in the cerebellum, myosin IIB1 is the dominant isoform, myosin IIB2 is weakly expressed, and the noninsertion type is not expressed (Itoh and Adelstein, 1995).

4.2.2 Differential Function of Myosin II Isoforms Like smooth muscle myosin II, activities of myosin IIA and myosin IIB are regulated by phosphorylation. Myosin II only becomes active when its RLCs are phosphorylated by myosin light chain kinase or Rho kinase. Myosin IIA is likely to have higher enzymatic activities than myosin IIB in vitro. In affinity purification of myosin IIA and myosin IIB, the speed of sliding of F‐actin along myosin is 3.3‐fold faster

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for myosin IIA than for myosin IIB, and Vmax of actin‐activated ATPase activity is 2.6‐fold higher for myosin IIA than for myosin IIB, although the affinities of myosin IIA and myosin IIB for F‐actin are almost equal (Kelley et al., 1996). However, it is unknown whether there are differences in enzymatic activities between myosin IIB isoforms.

4.2.3 Subcellular Localization of Myosin II Isoforms in Growth Cone In neuronal growth cones, myosin IIB shows spotted distribution throughout the cytoplasm, especially concentrated in the P‐domain along filopodia and the marginal zone between the P‐ and C‐domain (Rochlin et al., 1995). On the other hand, myosin IIA localizes in the C‐domain, but its expression level is much lower than that of myosin IIB (Rochlin et al., 1995). Evidence suggests that myosin IIA and myosin IIB play distinct roles in neurite dynamics. Knockout of myosin IIB with its antisense oligonucleotide inhibits neurite outgrowth in a differentiated neuroblastoma cell line (Wylie et al., 1998). A similar result is obtained with cultured neurons from myosin IIB knockout mice (Bridgman et al., 2001). However, knockout of myosin IIA with its antisense oligonucleotide weakens the adhesion of growth cones to the substrate (Wylie and Chantler, 2001) and inhibits LPA‐induced neurite retraction (Wylie and Chantler, 2003).

4.3 Myosin I Myosin I is a single‐headed motor composed of a HC (110–140kDa) and several LCs, all of which act as calmodulins in vertebrates. Myosin I is classified into four subclasses. The subclass 1 HC contains one IQ motif and one SH3 homology domain in its tail; human myosin Ic, human myosin Id, rat myr3, and lower eukaryote myosin I belong to this subclass. Subclass 2 HC contains 3–6 IQ motifs and can bind to the cell membrane via its tail; rat myr1, mouse MMIa and variety of brush border myosin I molecules are classified in this subclass. Subclass 3 HC contains three IQ motifs; mouse MMIb and rat myr2 are classified in this subclass. Subclass 4 HC contains two IQ motifs; mouse MMIg and rat myr4 are classified in this subclass. All four subclasses are expressed in the brain. Activities and regulatory modes of myosin I differ markedly among its subclasses. In the presence of mM‐order concentrations of Ca2þ, the ATPase activity of subclass 1 (myr3) decreases to one‐third of that in the absence of Ca2þ (Stoffler and Bahler, 1998). In contrast, the ATPase activity of subclass 3 (myosin Ib is increased twofold in Ca2þ concentrations of 1–10 mM (Barylko et al., 1992; Zhu et al., 1998). Activities of subclass 2 and subclass 4 have not been reported. However, in an in vitro motility assay, F‐actin slid along subclass 3 myosin Ib with a velocity of 0.33 mm/s in the absence of Ca2þ, but did not slide along subclass 3 myosin Ib in the presence of 1 mM Ca2þ (Zhu et al., 1998); these results contradict the results of ATPase measurements. At a low concentration of Ca2þ, subclass 3 myosin Ib appears to move along F‐actin with little consumption of ATP; in an in vitro motility assay, similar Ca2þ regulation was observed for subclass 2 (Williams, R., and Coluccio, L. M., 1994).

4.4 Myosin V Myosin V is composed of one pair of HCs (210 kDa), four pairs of calmodulin, two distinct pairs (23 and 17 kDa) of LCs, and an 8‐kDa dynein light chain associated with the tail domain (Cheney et al., 1993). Three isoforms, myosin Va, Vb, and Vc, have been found in humans, and myosin Va and Vb are expressed in the nervous system. Myosin Va is the most abundantly expressed and broadly distributed isoform (Mercer et al., 1991). Myosin Vb is localized in a restricted area including the hippocampus, dentate gyrus, and amygdala (Zhao et al., 1996). In neuronal growth cones, myosin Va colocalizes with 50–100‐nm vesicles, actin, and the plasma membrane (Evans et al., 1997). Disruption of myosin V causes inhibition of filopodia elongation in DRG neurons (Wang et al., 1996), although primary culture of superior cervical ganglion cells from myosin Va knockout mouse shows normal growth cone morphology (Evans et al., 1997).


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Unlike myosin II, myosin V has high affinity for F‐actin, even in the presence of ATP (De La Cruz et al., 1999; Trybus et al., 1999). Furthermore, myosin V does not detach from F‐actin during its catalytic cycle (Mehta et al., 1999); i.e., it is a ‘‘processive’’ motor. This processivity may allow a single myosin V molecule to function as a vesicle transporter along F‐actin tracks. The mechanism of processive movement of myosin V is controversial. Electron microscopic observations show that two myosin heads bind to F‐actin at a distance of 36 nm, which corresponds to the half‐helical pitch of F‐actin (Walker et al., 2000). Given this observation, Walker et al. have proposed that myosin V moves along F‐actin filaments ‘‘hand over hand’’ using two heads, and that at least one head is always attached to the F‐actin; this may be the source of the processivity of myosin V. However, if this is the case, single‐head myosin V would not show processivity, but it has been reported that single‐head myosin V has processivity (Tanaka et al., 2002). Further characterization of myosin V should clarify this issue.

4.5 Other Minor Myosins in Nervous System 4.5.1 Myosin III Myosin III is a single‐headed myosin with a N‐terminal kinase homology domain next to the head domain, two IQ motifs, and a C‐terminal tail domain. Two myosin III genes, myosin IIIA and myosin IIIB, have been found in vertebrates. Myosin IIIA is expressed in the retina and cochlea, and mutation of this gene causes progressive nonsyndromic hearing loss (Walsh et al., 2002). Baculovirus‐expressed myosin III has motor activity and kinase activity in vitro (Komaba et al., 2003), although the physiological roles of its kinase activity are unclear.

4.5.2 Myosin VI Myosin VI is a double‐headed myosin with a coiled‐coil structure between the neck domain and tail domain of its HC. Unlike all other known myosins, myosin VI is a minus end‐directed motor; i.e., it slides toward the minus (or pointed) end of F‐actin filaments (Wells et al., 1999). It localizes in inner‐ear hair cells (Hasson et al., 1997) and growth cones of cultured neurons (Suter et al., 2000). Mutation of the myosin VI gene is responsible for mouse Snell’s waltzer deafness, and evidence suggests that it affects the morphogenesis of hair cells (Avraham et al., 1995); however, its other physiological roles are unclear.

4.5.3 Myosin VII Myosin VII is a double‐headed myosin that localizes in inner‐ear hair cells (Hasson et al., 1997) and has been shown to be responsible for Usher 1B syndrome (Weil et al., 1996).

4.5.4 Myosin IX Myosin IX is a single‐headed myosin with four IQ motifs and a GTPase‐activating protein (GAP) domain in its heavy chain that is homologous to the GAPs of the Rho family (Chieregatti et al., 1998). It moves processively along F‐actin (Post et al., 2002).

4.5.5 Myosin X Myosin X is a double‐headed myosin containing three IQ motifs and a band 4.1/ezrin/radixin/moesin (FERM) domain in its HC (Berg et al., 2000). It localizes in dynamic actin structures such as lamellipodia, membrane ruffle, and filopodia (Berg et al., 2000). Myosin X is concentrated in the tips of filopodia and

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undergoes forward and rearward movement within filopodia (Berg and Cheney, 2002). Furthermore, overexpression of myosin X causes an increase in the number and length of filopodia suggesting that myosin X affects filopodia dynamics (Berg and Cheney, 2002). Myosin X also interacts with the cytoplasmic domain of b‐integrin via the FERM domain of myosin X heavy chains (Zhang et al., 2004).

5

Conclusion: Possible Roles of Actin, Actin‐Binding proteins, and Myosins in Living Cells

Thanks to innovations in imaging techniques, experimental data about the dynamics of actin, myosin, and actin‐binding proteins in neuronal cells has rapidly been accumulating in recent years. There has also recently been an increase in the rate at which data about these proteins has been obtained from in vitro reconstituted systems and analysis of specific gene disruption, as indicated from this chapter. The findings of these studies suggest harmonic interactions between actin, myosins, and actin‐binding proteins in neuronal cells. For example, most of the proteins presented in this chapter are expressed in neuronal growth cones. The available evidence suggests that they function as follows in growth cone dynamics. In the tips of filopodia, mDir accelerates the incorporation of actin. Fascin fastens to F‐actin to maintain its arrangement in tight bundles. Myosin IIB, V, X or other myosins translocate F‐actin bundles backward and cause retrograde flow of filopodial actin. Drebrin, tropomyosin, and caldesmon may loosen the F‐actin bundles, causing disassembly of the bundles at the border of the P‐domain and C‐ domain of the growth cone. Gelsolin, thymosin b and ADF/cofilin cause the depolymerization of F‐actin. In the tips of lamellipodia, N‐WASP coupled with the Arp2/3 complex accelerates actin polymerization; profilin promotes this process. Incorporated Arp2/3 complex causes branching of F‐actin filaments, resulting in formation of an F‐actin network. At the border of the P‐ and C‐domains, gelsolin, thymosin b and ADF/cofilin cause depolymerization of F‐actin. Myosin IIA and a‐actinin enhance the attachment of adhesion plaques. Myosin I and myosin V translocate membrane vesicles. In growth cones, these proteins may form a kind of community, with the cooperation of several proteins as a team in important functions. Their activity causes the dynamic motility of the growth cone (such as backward and forward movement), the cellular responses to a variety of guidance molecules, and the formation of neuronal networks.

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Transglutaminase

A. J. L. Cooper . S.‐Y. Kim

1

History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244

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Reactions Catalyzed by TGases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244

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The Human Family of TGases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 247

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General Physiological Roles of the TGases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 247

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Physiological Roles of TGases in Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249

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Possible Involvement of TGase in Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249 CAG‐Expansion Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249 Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 252 Other Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253

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TGases as Potential Therapeutic Targets in Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . 253

8

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254

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Abstract: Transglutaminases (TGases) catalyze the Ca2þ‐dependent cross‐linking of a glutaminyl (Q) residue in a protein or peptide substrate to a lysyl (K) residue in another protein or peptide substrate. These enzymes also catalyze the nucleophilic attack of amines, diamines, or polyamines on Q residues of protein or peptide substrates. TGases occur widely in nature. In rodents and humans, the TGase family is composed of nine members, eight of which are catalytically active. Human and mouse brain contain at least three TGase isoenzymes, namely TGase 1, TGase 2 (tTGase; so named because it is present in most mammalian tissues), and TGase 3. These enzymes have been implicated in some specialized neuronal functions, but generally their normal physiological roles in the central nervous system (CNS) are not well understood and have not been systematically studied. In contrast, the possible involvement of TGases in neurodegenerative diseases has been much more widely studied, especially within the last 5 years. As a result, this chapter contains more information on the roles of TGase in diseased brains than in healthy brains. Brain TGase activity and protein cross‐linking are increased in several neurodegenerative diseases, including Alzheimer’s disease (AD), Huntington’s disease (HD), Parkinson’s disease (PD), and supranuclear palsy (SNP). Such increased activity may at first be protective by removing damaged and unfolded protein, but with time the process may become pathological and contribute to the downward spiral in neurodegenerative diseases. If this hypothesis is confirmed, specific TGase inhibitors may be of therapeutic benefit in these diseases. List of Abbreviations: AD, Huntington disease; BDNF, brain derived neurotrophic factor; CBP, CRE binding protein; CNS, central nervous system; CSF, cerebrospinal fluid; CRE, cAMP response element; DPLA, dentatorubral-pallidoluysian atrophy; GGEL, N E-(γ-L-glutamyl)-L-lysine; HD, Huntington disease; htt, huntingtin; K, lysine; PD, Parkinson disease; PSP, progressive supranuclear palsy; Q, glutamine; Qn, polyglutamine; NRSE, neuronal restriction silencer element; SBMA, spinobulbar muscular atrophy (Kennedy’s disease); SCA, spinocerebellar ataxia; SNP, supranuclear palsy; TBP, TATA binding protein; TGase, transglutaminase

1

History

Heinrich Waelsch and colleagues were the first to describe a Ca2þ‐dependent enzymatic activity that incorporated amines into proteins with the concomitant formation of ammonia (Sarkar et al., 1957; Clarke et al., 1959). The authors termed this activity transglutaminase (TGase). TGase activity was found to be present in mammalian tissues including brain. Shortly after this discovery, Laszlo Lorand and colleagues discovered that plasma coagulation factor XIIIa possessed TGase activity (Lorand et al., 1962). (For a recent account of this early work see Lorand (2002).) Subsequently, TGase activity has been found in many organisms including bacteria, protozoa, plants, and vertebrates (Serafini‐Fracassini et al., 1995; Griffin et al., 2002; Wada et al., 2002; Lorand and Graham, 2003; Zotzel et al., 2003; Mea et al., 2004).

2

Reactions Catalyzed by TGases

TGases catalyze the attack of a suitable nucleophile (acyl acceptor) at the carboxamide group of a glutaminyl residue (Q) of a protein or peptide substrate (acyl donor) (> Figure 7-1). The nucleophile may be the E amino group of a lysyl residue (K) in a protein or polypeptide (reaction 1), an amine, diamine, or polyamine (reaction 2), water (reaction 3), or polyamine already bound at a glutaminyl residue (reaction 4) (Folk, 1983). In addition, it has been reported that TGase 1 can catalyze the esterification of carboxamide residues of the skin protein involucrin with the o‐hydroxy group of an artificial o‐hydroxy ceramide (Nemes et al., 1999) (schematically depicted in reaction 5). The reaction with water (a hydrolysis reaction; reaction 3) results in the deamidation of Q residues to glutamate (E) residues. Deamidation alters the charge on the protein or peptide, and under certain circumstances this may result in an immunological response. For example, TGase‐2‐catalyzed deamidation of a Q‐containing 33‐mer peptide derived from gliadin (a protein present in wheat, oats, and rye) is apparently responsible, at least in part, for the

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. Figure 7-1 Reactions catalyzed by various Ca2þ‐dependent transglutaminases (TGases). Protein and peptide substrates are depicted as rectangles and ovals, respectively. Only the glutaminyl and lysyl residue of the protein and peptide substrates are depicted. The carboxamide group of the glutaminyl residue is the acyl donor. The acyl acceptor (attacking nucleophile) may be: (1) a lysyl residue of a protein or peptide; (2) an amine, diamine or polyamine; (3) water; (4) a polyamine already attached to a glutaminyl residue; or (5) an ester. Quantitatively, the more important reactions are 1 and 2. Hydrolysis occurs only under conditions of limiting amine substrate. Esterification has only been shown to be of importance with TGase 1. In addition, it has been known for about 10 years that TGase 2 binds and hydrolyzes GTP. Recently, TGase 3, TGase 4, and TGase 5 have also been shown to bind and hydrolyze GTP. In addition, TGase 2 has now been shown to catalyze a kinase reaction (see text). Adapted from Lorand (2002)

immunological intolerance and damage to the proximal small intestine wall in celiac disease (celiac sprue) (Reif and Lerner, 2004). A protein (Gha1) initially identified as being involved in receptor signaling function was later identified as TGase 2 by Graham and coworkers (Nakaoka et al., 1994; Iismaa et al., 2000). Binding of GTP elevates the Ca2þ requirement and inhibits the transamidating activity (Liu et al., 2002). GTP is hydrolyzed by TGase 2 (> Figure 7-1, reaction 6). TGase 2 is therefore a bifunctional enzyme that catalyzes two distinct enzymatic reactions. The two reactions are catalyzed at distinct sites on the enzyme surface. Subsequently, three other TGases (TGase 3, TGase 4, and TGase 5) were found to bind and hydrolyze GTP (Mariniello et al., 2003; Ahvazi et al., 2004; Candi et al., 2004). TGase 2/Gha1 is thought to be a signal transducer of

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a1‐adrenergic receptor‐induced smooth cell proliferation (Dupuis et al., 2004). The biological functions of the GTP binding and GTPase reactions of TGase 3, TGase 4, and TGase 5 are not certain. TGase 2 has recently been shown to possess kinase activity and to catalyze phosphorylation of insulin‐ like growth factor‐binding protein‐3 (Mishra and Murphy, 2004). Evidence was presented that TGase 2 is the major kinase responsible for the phosphorylation of insulin‐like growth factor‐binding protein‐3 in breast cancer cells (Mishra and Murphy, 2004). TGase 2 has also recently been found to possess protein disulfide isomerase activity (Hasegawa et al., 2003). Evidently, the potential biological roles of the TGase family are continuing to expand. Here, we concentrate on the well‐defined transamidating ability of the TGases. Some crystal structures are available for TGase 2, factor XIIIa, sea bream TGase, and TGase 3. (see Iismaa et al., (2003) and references cited therein.) Apparently, in the absence of Ca2þ, a conserved tryptophan (W) residue obstructs the active site, but in the presence of Ca2þ the W residue moves, thereby opening a channel. This opening allows the side arm of a Q residue of the protein or peptide substrate to be threaded into the channel in such a way that the carboxamide comes in contact with an active site cysteine (Chica et al., 2004). Asparagine residues are not substrates, which suggests that the side arm of the Q residue substrate is fully extended in the active site channel. All members of the classical eukaryotic TGase family investigated thus far possess an active site that is similar to that of the papain‐like family of cysteine proteases. The active site possesses a Cys‐His‐Asp or Cys‐His‐Asn triad, in which the Cys and His residues probably form a thiolate–imidazolium ion pair. The TGases catalyze a Ping‐Pong reaction that involves a cycle of acylation at the active site cysteine residue followed by deacylation (> Figure 7-2). The Q‐containing substrate forms a Michaelis complex, which

. Figure 7-2 TGase‐catalyzed Ping‐Pong mechanism. The large horizontal arrow represents the progress of the reaction and depicts one turnover at the active site. The incoming Q substrate (acyl donor) forms a Michaelis complex with the thiolate anion of an active site cysteine residue (E‐S). Attack of the thiolate at the carboxamide of the Q residue generates the first tetrahedral (oxyanion) intermediate. This intermediate decomposes to an active site‐ attached thioester (acyl‐enzyme intermediate) with the concomitant release of ammonia. Next, the amine substrate (R’NH2, acyl acceptor) enters the active site and forms a Michaelis complex with the thioester intermediate. Nucleophilic attack by the N of the amine at the carbonyl carbon of the thioester generates the second tetrahedral (oxyanion) intermediate. This intermediate spontaneously decomposes to free enzyme (E‐S) and cross‐linked product. Cross‐linking is through an N e‐(g‐L‐glutamyl)‐L‐lysine (GGEL) linkage or g‐L‐glutamylamine linkage

enables the thiolate of the cysteine residue to attack the carboxamide of the Q residue forming a tetrahedral (oxyanion) intermediate at the active site. The tetrahedral intermediate then breaks down to generate a thioester (acyl) intermediate and ammonia. This enzyme acylation reaction is rate limiting. In the next step, the second substrate (acyl acceptor), whether it is a small‐Mr amine/diamine/polyamine or a K residue

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of a protein or peptide, forms a second Michaelis complex. Nucleophilic attack by the nitrogen generates a second tetrahedral (oxyanion) intermediate, which breaks down to product and free enzyme (> Figure 7-2). In TGase 2, the transition state intermediates are stabilized by a W and tyrosine (Y) residue in the active site (Iismaa et al., 2003; Chica et al., 2004). The Y residue also stabilizes the oxyanion intermediates. Residues adjacent to the Q residue play an important role in enabling the Q residue to be threaded into the active site (Chica et al., 2004).

3

The Human Family of TGases

The human and mouse genomes each encode nine closely related proteins that belong to the TGase family. Eight of these are active in catalyzing transamidating reactions (TGases 1–7; factor XIIIa) and one is inactive (erythrocyte band 4.2). These proteins are variously expressed in different organs and tissues (Grenard et al., 2001). A list of human TGases is given in > Table 7-1. It should be noted that TGase 2 is not activated by proteolytic processing, but other TGases (e.g., TGase 1, TGase 3) are synthesized initially as zymogens . Table 7-1 Proteins in the TGase family and some of their major biological functions where known TGase Factor XIIIa TGase 1 (keratinocyte TGase, kTGase) TGase 2 (tissue TGase, tTGase, cTGase) TGase 3 (epidermal TGase, eTGase) TGase 4 (prostate TGase, pTGase) TGase 5 (TGase X) TGase 6 (TGase Y) TGase 7 (TGase Z) Erythrocyte band 4.2

Biological function Blood clotting Skin differentiation Apoptosis, cell adhesion, signal transduction, protein quality control Hair follicle differentiation, epidermis maturation Suppression of sperm immunogenicity in the female genital tract, formation of genital plug in rodents Epidermal differentiation Unknown function; possibly epidermal differentiation Unknown function; possibly epidermal differentiation Component of erythrocyte cell wall

Synonyms are presented in parenthesis. Erythrocyte band 4.2 clearly belongs to the TGase gene family, but does not contain an active site cysteine residue. Therefore, this protein cannot catalyze transglutamination reactions. This protein is a structural component of red blood cells. Mutations in this protein cause rare red cell membrane disorders (Ialascon et al., 2003)

that are activated by proteolytic ‘‘nicking’’. Nicking activates these TGases 100–1000‐fold (Kim et al., 1994). Factor XIII is converted to the active TGase form (factor XIIIa) through proteolytic nicking catalyzed by thrombin. A generalized structure of human TGase 2 showing various binding and structural domains is shown in > Figure 7-3.

4

General Physiological Roles of the TGases

The major known physiological function of these enzymes is to catalyze the formation of an isopeptide N E‐(g‐L‐glutamyl)‐L‐lysine (GGEL) linkage between the carboxamide moiety of protein Q residue and the E‐amino group of a protein K residue (> Figure 7-1; reaction 1). Factor XIIIa catalyzes the formation of hard clot and is essential in preventing excessive bleeding after injury. Mutations in this protein cause protracted bleeding (Mikkola and Palotie, 1996). TGase 1 catalyzes cross‐linking of proteins that comprise the cell envelope, a flexible insoluble barrier that lines the outer surface of fully differentiated cells of squamous epithelia, thereby protecting the body against mechanical and chemical damage, and against dehydration. Mutations in the TGase 1 gene that result in loss‐of‐function, mutations in TGase 1 that

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. Figure 7-3 Schematic diagram of human transglutaminase 2 (TGase 2). The active enzyme is a monomer with a Mr of 78,000. The numbers refer to amino acid residues distal to the N terminus. The main binding sites for fibronectin, phospholipase C, Ca2þ, and GTP/ATP are shown. The active site for the GTP/ATPase and the active site cysteine involved in the transamidating activity are also shown. The residues involved in protein disulfide isomerase activity are not known, but do not appear to involve the Ca2þ‐binding region or the transglutaminase active site cysteine (Hasegawa et al., 2003). The residues responsible for the insulin‐like growth factor‐ binding protein‐3 kinase activity are also not known (Mishra and Murphy, 2004)

impede proteolytic activation, and mutations in other enzymes that result in altered catalytic properties of TGase 1 lead to a family of severe skin diseases known as ichthyoses (reviewed by Kim et al., 2002a). TGase 3 is involved in the formation of the cell envelope. Various soluble cell envelope proteins are cross‐linked by cytosolic TGase 3 into oligomers, which are then transported to the cell periphery where they are cross‐ linked to the growing cell envelope by the membrane‐anchored TGase 1 (Kim et al., 2002a and references cited therein). There are no known human diseases caused by aberrant TGase 3 activity, and TGase 3 knockout leads to very early embryonic lethality in mice (L. M. Milstone, personal communication). TGase 4 is largely confined to the prostate. The enzyme appears to be involved in suppression of sperm immunogenicity in the female genital tract. In rodents, TGase 4 is also involved in the formation of the vaginal plug. TGases 5–7 have been relatively little studied, but may be involved in specialized epidermal differentiation. Several Q residues of histones H2A, H2B, H3, and H4 under appropriate conditions in isolated core particles/nucleosomes are acyl donor substrates of TGases (Ballestar et al., 2001; Kim et al., 2002b). Substrate accessibility depends on ionic strength and DNA dissociation. We have shown that the K‐rich linker histone (histone H1) is an excellent acyl acceptor of TGase 2 in vitro (Cooper et al., 2000). It is well‐known that nuclear histones may be altered by acylation and methylation. Such alterations play a role in gene expression (Fischle et al., 2003). It is possible that the action of TGase on histones adds yet another layer in the complexity of gene regulation (Cooper et al., 2000; Kim et al., 2001, 2002b). The most thoroughly studied member of the TGase family is TGase 2. The enzyme has been implicated in a number of pathological conditions including fibrosis, atherosclerosis, celiac disease, cataractogenesis, neurodegenerative disease (see below), and cancer metastasis (see, for example, Aeschlimann and Thomazy, 2000; Kim et al., 2002a; Bailey et al., 2004; Shin et al., 2004; and references cited therein). TGase 2 has also been implicated in a number of natural biological functions including cellular proliferation, cellular differentiation, apoptosis, cell adhesion, wound healing, and signal transduction. Mice containing

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a TGase 2 knockout (unlike the situation with TGase 1 and TGase 3 knockouts) exhibit no overt, major phenotypic abnormalities (De Laurenzi and Melino, 2001). The TGase 2/ mice are, however, more susceptible to liver injury (Nardacci et al., 2003) than their TGase 2þ/þ littermates. Abnormalities in thymocyte viability and fibroblast adhesion are also present in the TGase 2/ mice (Nanda et al., 2001). Possibly, other TGases compensate for most of the lost functions of TGase 2 in vivo in the TGase 2/ mice.

5

Physiological Roles of TGases in Nervous Tissue

Kim et al. (1999) showed that human brain contains mRNA and protein for TGases 1, 2, and 3. mRNA for TGase 5 and TGase 7 are present at relatively low levels in the human brain (Grenard et al., 2001), but the level of expression of the corresponding proteins is uncertain. TGase 2 is the most active TGase isoform in normal brain (as assessed by a commonly used assay that measures incorporation of radiolabeled polyamines into dimethyl casein or succinylated casein). Some reports show TGase 2 to be present in cultured astrocytes (Reichelt and Poulsen, 1992; Campisi et al., 2003). Other studies showed TGase 2 to be present in hippocampal neurons (Appelt et al., 1996), in neurons in the spinal cord (Perry and Haynes, 1993]) and in cultured cerebellar granule cells (Perry et al., 1995). TGase activity is present in many regions of the rat brain. Specific activities are comparable in all these regions (Gilad and Varon, 1985). Activity is also present in all subcellular fractions studied, although the highest activity is in the cytosolic compartment. Activity is present on the surface of synaptosomes (Gilad and Varon, 1985). In skeletal muscle, TGase 2 is most concentrated and active at the motor endplate. In the brain, TGases have been proposed to play a role in suppression of vesicular catecholamine release (Pastusko et al., 1986), neuronal differentiation (Macchione and Seeds, 1986; Hand et al., 1993; Tucholski et al., 2001), neuronal apoptosis (Piacentini et al., 1992; Melino et al., 1994), neuronal outgrowth, guidance and sustainment of shape (Hand et al., 1993), and long‐term potentiation (Friedrich et al., 1991).

6

Possible Involvement of TGase in Neurodegenerative Diseases

6.1 CAG‐Expansion Diseases At least ten inherited neurodegenerative diseases are caused by a (CAG)n expansion in the affected gene (Maryuma et al., 2002; Taylor et al., 2002; Zoghbi and Orr, 2002) (> Table 7-2). The most common of these diseases is HD. Others are dentatorubral‐pallidoluysian atrophy (DRPLA), Kennedy’s disease (spinobulbar muscular atrophy, SBMA), and seven forms of spinocerebellar ataxia (SCA1, SCA2, SCA3 (Machado– Joseph disease), SCA6, SCA7, SCA12, and SCA17). The CAG‐expansion diseases are usually adult‐onset with progressive worsening of symptoms and eventual death. The genes responsible appear to be unrelated except for the fact that each possesses a CAG expansion. In every case, except SCA12, the mutation is in a coding region. The mutated protein possesses an expanded Qn domain and appears to be normally expressed. Many of these diseases are characterized by insoluble protein aggregates in the affected areas. The aggregates contain the mutated protein. The CAG‐expansions (except in the case of SCA12) are widely thought to confer a pathological gain‐of‐function to the expressed protein, although in some cases a pathological loss‐of‐function may also contribute. Generally, the longer the repeat the earlier the onset and the more rapid and severe the disease progression. This phenomenon is particularly well documented in HD. The possibility has been considered that the protein aggregates in brains from patients with HD are toxic (Bates, 2003). However, this notion is controversial. Some authors suggest that the aggregates are neutral or even beneficial (Saudou et al., 1998; Kuemmerle et al., 1999; see the discussion by Michalik and Broeckhoven (2003)). In order to gain insights into the pathological mechanisms associated with the CAG‐ expansion diseases, it is important to understand the origin of the protein aggregates in CAG‐expansion diseases and whether or not the deposits contribute to the disease process.

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. Table 7-2 Current list of CAG‐expansion diseasesa

Disease

Sites of neuropathology

Normal

Disease

Gene product (intracellular localization of aggregates)

Corea major or Huntington’s disease (HD) Spinocerebellar ataxia type 1 (SCA1) Spinocerebellar ataxia type 2 (SCA2) Spinocerebellar ataxia type 3 (SCA3) or Machado–Joseph disease (MJD) Spinocerebellar ataxia type 6 (SCA6)b,c Spinocerebellar ataxia type 7 (SCA7) Spinocerebellar ataxia type 12 (SCA12)d

Striatum (medium spiny neurons and cortex in late stage) Cerebellar cortex (Purkinje cells), dentate nucleus, and brain stem Cerebellum, pontine nuclei, substantia nigra Substantia nigra, globus pallidus, pontine nucleus, cerebellar cortex Cerebellar and mild brainstem atrophy Photoreceptor and bipolar cells, cerebellar cortex, brainstem Cortical, cerebellar atrophy

6–35

36–121

Huntingtin (n, c)

6–39

40–81

Ataxin‐1 (n, c)

15–29

35–64

Ataxin‐2 (c)

13–42

61–84

Ataxin‐3 (c)

4–18

21–30

7–17

37–130

Calcium channel subunit (a1A) (m) Ataxin‐7 (n)

7–32

41–78

Spinocerebellar ataxia type 17 (SCA 17) Spinobulbar muscular atrophy (SBMA) or Kennedy’s disease

Gliosis and neuronal loss in the Purkinje cell layer Motor neurons (anterior horn cells, bulbar neurons) and dorsal root ganglia Globus pallidus, dentato‐rubral, and subthalamic nucleus

29–42

46–63

11–34

40–62

7–35

49–88

CAG repeat number

Dentatorubral‐pallidoluysian atrophy (DRPLA)

Brain specific regulatory subunit of protein phosphatase PP2A TATA‐binding protein (TBP) (n) Androgen receptor (n, c) Atrophin (n, c)

a

Adapted from Gentile and Cooper (2004). Cellular localization of aggregates: c, cytosolic; m, transmembrane; n, nuclear The lowest pathological range of CAG repeats among the CAG‐expansion diseases occurs in SCA 6, and comparison with other mutations in the same gene suggests that expanded Qn domains in SCA6 affect the normal function of the calcium channel i.e., there is a toxic loss‐of‐function. In other cases, increased Qn domains result in a toxic gain‐of‐function or a mixture of the two (see text) c Qn expansions may occur in the potassium channel protein in some familial forms of schizophrenia, but the notion remains controversial d In all cases except SCA 12, the mutation occurs in the coding region of the gene. In SCA12, the CAG expansion occurs in the untranslated region at the 5’‐end of the PPP2R2B gene. The toxicity may result from ‘‘overexpression’’ of the brain specific regulatory (PR55b) subunit of protein phosphatase PP2A (Holmes et al., 2003) b

Previously, two main theories were posited concerning the origin of the insoluble brain deposits in CAG expansion diseases—the polar zipper hypothesis and the TGase hypothesis (reviewed by Cooper et al., 2002). The polar zipper theory suggests that Qn domains interact noncovalently to produce insoluble, ordered structures that are more prominent with the larger Qn domains. In contrast, deposits formed by TGase action have covalent cross linkages and have nonordered structures (Karpuj et al., 1999). Iuchi et al. (2003) showed that formic acid‐resistant oligomeric proteins are present in the cerebral cortex (heavily affected region) of patients suffering from HD, but not in the cerebellum (less affected region). Resistance to formic acid suggests the occurrence of proteins cross‐linked by TGases (Iuchi et al., 2003). Several studies have shown that TGase 2‐catalyzed cross‐linking renders many proteins insoluble (Kahlem et al., 1998).

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However, TGase 2‐catalyzed cross‐linking can promote solubilization of proteins in some cases (Lai et al., 2004). Moreover, Mastroberardino et al., (2002) showed a 30% increase in brain intranuclear inclusions in R6/1 TGase 2/ knockout mice compared to the R6/1 TGase 2þ/þ mice together with a 90% reduction in GGEL linkages. Evidently, which process—polar zippers or TGase‐catalyzed covalent bond formation— predominates in aggregate formation may vary depending, for example, on the size of the Qn insert, levels of mutated protein, TGase, Ca2þ, GTP, and the nature of the Q/K substrates. The toxic gain‐of‐function associated with CAG‐expansion diseases may be due to an increase in TGase activity (Green et al., 1993), whether or not the resulting cross‐linked proteins contribute to insoluble protein aggregates. There is considerable indirect evidence to support this hypothesis. For example, two groups have reported that total TGase activity is increased in brains of patients with HD (Karpuj et al., 1999; Lesort et al., 1999). Additionally, Lesort et al., (1999) reported a stage‐dependent increase in TGase 2 message and protein in brains of patients with HD. An influx of intracellular Ca2þ together with higher TGase activity, favorable substrate (i.e., elongated Qn domain), and perhaps lowered GTP levels suggest that a disinhibition of TGase in brains of patients with HD leads to higher and potentially harmful TGase activity (Lesort et al., 2000). Qn domains, especially those of a pathological length, are excellent TGase 2 substrates (Kahlem et al., 1996, 1998; Cooper et al., 1997; Gentile et al., 1998; Lesort et al., 1999). Calmodulin was shown to coimmunoprecipitate with TGase 2 and huntingtin (htt; the mutated protein in HD) in cells transfected with myc‐tagged N‐terminal htt fragments containing 148 Q repeats and TGase 2, but not in cells containing a construct with 18 Q repeats. A calmodulin inhibitor lessened the number of cross‐ links and aggregates (Zainelli et al., 2004). It was suggested that calmodulin regulates TGase 2 activity in brain (Zainelli et al., 2004). TGase 2‐like immunoreactivity was shown to colocalize with ubiquitinated intranuclear inclusions in brains of patients with DRPLA (Sato et al., 2002). Two studies have shown that the in vitro TGase inhibitor cystamine prolongs the lives of R6/2 mice (a transgenic model of HD), lessens weight loss, and improves behavioral functions (Dedeoglu et al., 2002; Karpuj et al., 2002). In one of these studies, brain aggregates were decreased (Dedeoglu et al., 2002). A particularly important study is that of Mastroberardino et al. (2002) who reported a reduction in neuronal cell death, improved behavior, and prolonged survival in R6/1 [HD] TGase 2/ (knockout) mice compared to R6/1 TGase 2þ/þ mice with intact TGase 2. How might increased TGase activity contribute to neurodegeneration in CAG‐expansion diseases? We have proposed that the answer may lie with the increasing ability of Qn repeats to act as TGase substrates as chain length increases. Such a theory can explain: (1) the relatively sharp demarcation between nonpathological (n ¼ 35) and pathological (n ¼ 40) Qn domains in HD and most other CAG‐expansion diseases; and (2) the decreasing age of onset and increased severity as n increases beyond 40 (Cooper et al., 1999, 2002). It is well known that brains afflicted with HD are under oxidative stress (Browne and Beal, 2004), and oxidative stress has been shown to promote increased TGase 2 expression (Shin et al., 2004). Oxidative stress also increases the incidence of damaged proteins. Inasmuch as exposed Q residues are more likely to be TGase substrates than buried residues, damaged/unfolded proteins may be better substrates for TGase 2 than undamaged proteins. Thus, increased TGase activity may be beneficial at first by removing or altering the properties of damaged proteins. However, this may become a double‐edged sword if the increased TGase activity eventually leads to loss of proteins or peptides necessary for normal brain function. We have suggested that excessive cross‐linking of Qn repeats may eventually cause the proteasome‐ degrading machinery to malfunction (Cooper et al., 2002). Recently, Mandrusiak et al. (2003) showed that the N terminus of the SBMA protein (the androgen receptor, AR) that contains the Qn repeat is an excellent Q and K substrate of TGase 2, forming cross‐linked polymers. HEK GFPu‐1 cells expressing TGase 2 and mutated AR exhibited proteasome dysfunction. Moreover, GGEL cross‐links were detected immunohistochemically in the aggregates in the brains of SBMA transgenic mice, but not in the controls (Mandrusiak et al., 2003). The authors suggested that cross‐linked AR obstructs the proteasome pore, contributing to SBMA pathogenesis. Many transcription factors such as CRE‐binding protein (CBP) and TATA‐binding protein (TBP) contain Qn domains (Schaffar et al., 2004), which may assist in the assembly of the transcriptosome. Qn‐containing transcription factors are theoretically capable of forming polar zippers with the Qn domains of the proteins mutated in the CAG‐expansion diseases. In the case of HD, N‐terminal fragments of

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mutated htt (containing the expanded Qn domain) accumulate in the nucleus (Zainelli et al., 2003). Moreover, nuclear‐targeting of mutant htt fragments produces HD‐like phenotypes in HD transgenic mice (Schilling et al., 2004). Thus, it is possible that one of the toxic effects of expanded Qn domains is the alteration of transcription factor interactions (Schaffar et al., 2004). It has been shown that mutant htt binds to CBP and p53. The latter regulates transcription of various mitochondrial proteins, which may underlie in part the mitochondrial abnormalities in HD (Sawa, 2001). Wild type but not mutant htt stimulates transcription of brain‐derived neurotrophic factor (BDNF), and neuronal restrictive silencer element (NRSE) is the target of wild‐type htt activity on the BDNF promoter II (Zuccato et al., 2003). Recently, it was shown that mutant htt in a mouse model of HD facilitates CRE‐dependent transcription (Obrietan and Hoyt, 2004). Thus, mutated htt may cause either increases (Obrietan and Hoyt, 2004) or decreases (Zuccato et al., 2003) in transcriptional regulation. Such alterations are likely to contribute to the pathological response in HD and other CAG‐expansion diseases. Interactions between mutant htt and transcription factors may occur not only by polar zipper interactions, but also by covalent alterations. TGase activity is increased in the nuclei of HD brain cells (Karpuj et al., 1999; Lesort et al., 1999). Moreover, acyl acceptors (polyamines and possibly histones) and increased levels of a ‘‘good’’ acyl substrate (i.e., htt fragment with an expanded Qn domain) are present in these cell nuclei. These findings raise the possibility that htt fragments and transcription factors may be covalently ‘‘glued’’ together by TGases in the nuclei of cells from patients with HD. In this regard, it is interesting to note that one of the CAG‐expansion diseases (SCA17) is caused by an expansion of a transcription factor (TBP) (> Table 7-2), leading to aberrant gene expression. It is also theoretically possible that increased TGase‐catalyzed cross‐linking of histones may also lead to disrupted gene function in brains of patients with HD(Cooper et al., 2002). In addition, it has been shown that lymphoblast mitochondria isolated from patients with HD and mitochondria isolated from the brains of HD transgenic mice have a lower mitochondrial potential and depolarize at lower Ca2þ loads than do mitochondria from controls (Panov et al., 2002). These effects can be duplicated with normal mitochondria exposed to proteins containing mutant htt (Panov et al., 2003). On the other hand, TGase 2 overexpression sensitizes a neuronal cell line to apoptosis by increasing mitochondrial membrane potential and cellular oxidative stress (Piacentini et al., 2002). Moreover, in a neural cell model, impaired mitochondrial function led to increased TGase activity in situ (Lesort et al., 2000). Thus, oxidative stress could lead to increased TGase 2 activity, which in turn could lead to mitochondrial damage and further oxidative stress‐increased TGase activity, resulting in an inexorable downward spiral. The roles of aberrant TGase activity, mutant protein, and altered transcription factors on the properties of brain mitochondria in CAG‐expansion diseases need to be further investigated.

6.2 Alzheimer’s Disease The defining characteristic of brains of patients with AD is the presence of plaques and tangles. Some evidence suggests that TGases contribute to the formation of these insoluble structures. Selkoe et al. (1981, 1982) showed that brain TGases catalyze the formation of high‐Mr polymers with cytoskeletal elements in vitro and suggested that TGase‐catalyzed reactions might contribute to paired helical formation (PHF) in the tangles present in brains of patients with AD. Subsequently, it was shown in the brains of patients with AD that (1) TGase 2 colocalizes with tangles (Miller and Anderton, 1986; Johnson et al., 1997); (2) GGEL linkages can be detected immunohistochemically in the neurofibrillary tangles (Norlund et al., 1999; Citron et al., 2002; Singer et al., 2002); (3) TGase 2 can be detected immunohistochemically in the plaques and in isolated amyloid core plaques (Zhang et al., 1998); and (4) the number of GGEL isopeptide linkages in insoluble protein is greatly increased relative to those of control brains (Kim et al., 1999). Moreover, b‐amyloid (Ab), a component of AD plaques, is an excellent substrate of TGase 2 in vitro (Ikura et al., 1993; Dudek and Johnson, 1994; Ho et al., 1994; Rasmussen et al., 1994), as are tau (the main component of PHFs) (Miller and Anderton, 1986, Dudek and Johnson, 1993; Miller and Johnson, 1995; Appelt and Balin, 1997; Murthy et al., 1998) and the non‐Ab component derived from a‐synuclein (NAC) of AD plaques (Jensen et al., 1995). The Ab polymers resulting from TGase‐2‐catalyzed cross‐linking of Ab and the insoluble

Transglutaminase

7

filamentous products resulting from cross‐linking of tau strongly resemble polymeric products that can be isolated from the brains of patients with AD (Jensen et al., 1995; Appelt and Balin, 1997). Additional findings implicate aberrant TGase activity in brains of patients with AD. For example, in patients with AD, TGase 1 and 2 proteins are elevated in the brain (Kim et al., 1999), TGase 2 protein is elevated in the CSF (Bonelli et al., 2002), total TGase activity is increased in the brain (Johnson et al., 1997; Kim et al., 1999), and free GGEL is elevated in the CSF (Nemes et al., 2001). The increase in free GGEL in CSF of AD patients may be due to apoptotic turnover involving TGase 2 (Nemes et al., 2001). It is also possible that TGase 2 and other TGases generate an insult not directly related to apoptosis. Whatever the primary event in AD, a series of events appear to be set in motion that leads to greatly increased TGase activity in affected brain. This increased activity may at first be beneficial by removing misfolded or damaged proteins, but with time the process may remove essential proteins and peptides (perhaps including components of the proteasome), and thereby become detrimental. If this is the case, even though increased TGase activity is not the primary cause of the disease, an inhibitor of brain TGases may have therapeutic benefit in AD (and HD)—at least at appropriate stages of the disease.

6.3 Other Neurodegenerative Diseases a‐Synuclein is present in Lewy bodies in patients with PD (Arima et al., 1998) and in neuritic plaques in brains of patients with AD (Wirths et al., 2000). As noted above, a‐synuclein is a TGase 2 substrate. Moreover, GGEL cross‐links have been detected immunohistochemically in the Lewy bodies from brains of patients with PD (Citron et al., 2002). TGase 2 catalyzes the conversion of a‐synuclein into high‐Mr polymers (Junn et al., 2003). In a cellular model of PD, overexpression of a‐synuclein and TGase 2 resulted in the formation of detergent‐insoluble aggregates. Immunocytochemical studies revealed the presence of a‐synuclein‐positive cytosolic inclusions in 8% of the TGase 2‐expressing cells. Aggregate formation was increased by a calcium ionophore and abolished by cystamine (Junn et al., 2003). Neurofibrillary tangles occur in affected regions in the brains of patients with progressive supranuclear palsy (PSP), and tau has been identified as a major component of detergent‐insoluble proteins in affected brains (Flament et al., 1991). In brains of patients with PSP, the tau protein in the tangles has been shown immunohistochemically to contain GGEL cross‐links (Zemaitaitis et al., 2000). Overall TGase activity, TGase 1 mRNA and protein, TGase 2 mRNA (including a shortened form) are elevated in selected, damaged regions in the brains of patients with PSP (Zemaitaitis et al., 2003). Tissues with more TGase activity had more neurofibrillary tangles (Zemaitaitis et al., 2003). Increased expression of TGase 2 has been noted in damaged tissue in animal models of stroke (Ientile et al., 2004; Tolentino et al., 2004), traumatic brain injury (Tolentino et al., 2002), and spinal cord injury (Citron et al., 2002). The injury to nervous tissue promotes the appearance of a shortened (S) form of TGase 2 (Festoff et al., 2002). The importance of the S form in the pathological process is not clear. TGase activity is increased in the nuclei of cultured astroglial cells in response to excitotoxic levels of glutamate (Campisi et al., 2003). Finally, TGase activity is increased in the brains of senescent rats (Park et al., 1999; Kim et al., 2001). The authors suggested that TGase 2 might be a senescence marker. A note of caution: Questions have been raised regarding the specificity of commercially available TGase 2 and GGEL antibodies (Bailey et al., 2004; Johnson and LeShoure, 2004). Although there is overwhelming evidence for increased TGase activity and GGEL formation in diseased brains, caution may be required in interpreting some of the papers quoted here in which TGase 2 and GGEL were detected using commercial antibodies.

7

TGases as Potential Therapeutic Targets in Neurodegenerative Diseases

As mentioned above, two groups have shown that the in vitro TGase inhibitor cystamine is beneficial in a mouse model of HD (Dedeoglu et al., 2002; Karpuj et al., 2002). However, the results do not prove involvement of TGases in HD. It is most likely that cystamine is reduced to cysteamine in vivo. Cysteamine

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is a potent antioxidant and has been used as a radioprotectant in cancer therapy (Duffel et al., 1987). Inasmuch as the brains of patients with HD are under oxidative stress, it is conceivable that the beneficial effects of cystamine and cysteamine are related to their antioxidant properties rather than to their inhibition of TGase(s). However, in cell models of CAG‐expansion diseases (Mandrusiak et al., 2003) and PD (Junn et al., 2003), cystamine treatment lessens or abolishes TGase‐induced aggregate formation. This finding suggests that in certain cell models of neurodegeneration, cystamine does indeed interfere with TGase‐ catalyzed protein cross‐linking. Nevertheless, it will be important to determine whether compounds that are not antioxidants and which are specific TGase inhibitors will be of benefit in animal models of neurodegenerative diseases (Gentile and Cooper, 2004). It is important that such compounds discriminate between the brain TGases and factor XIIIa in order to prevent bleeding as an unacceptable side effect. This is particularly important in considering the use of such compounds to treat human diseases.

8

Conclusions

Although TGases have been implicated in a number of important functions in the CNS, the roles need to be better defined. On the other hand, considerable evidence suggests that many neurodegenerative diseases are associated with aberrant TGase activity. This process may at first be beneficial, but may become deleterious with time, contributing to the inexorable downward spiral so often associated with neurodegenerative disease. Some preliminary evidence suggests that TGase inhibitors may be beneficial, but a new generation of specific inhibitors is needed to prove the involvement of TGases in neurodegenerative disease. If these inhibitors prove successful in cell and animal models of neurodegenerative diseases then we can look forward to a new approach to the treatment of these dreadful diseases.

Acknowledgments Work from our laboratory mentioned here was supported in part by NIH grant 1PO1AG14930.

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Transglutaminase Candi E, Paradisis A, Terrinoni A, Pietroni V, Oddi S, et al. 2004. Transglutaminase 5 is regulated by guanine/adenine nucleotides. Biochem J 381: 313-319. Chica RA, Gagnon P, Keillor JW, Pelletier JN. 2004. Tissue transglutaminase acylation: Proposed role of conserved active site Tyr and Trp residues revealed by molecular modeling of peptide substrate binding. Protein Sci 13: 979-991. Citron BA, Suo Z, Santa Cruz K, Davies PJ, Qin F, et al. 2002. Protein cross‐linking, tissue transglutaminase, alternative splicing, and neurodegeneration. Neurochem Int 40: 69-78. Clarke DD, Mycek MJ, Neidle A, Waelsch H. 1959. The incorporation of amines into proteins. Arch Biochem Biophys 79: 338-354. Cooper AJL, Jeitner TM, Gentile V, Blass JP. 2002. Cross‐ linking of polyglutamine domains catalyzed by tissue transglutaminase is greatly favored with pathological length repeats: Does transglutaminase activity play a role in (CAG)n/Qn‐expansion diseases? Neurochem Int 40: 53-67. Cooper AJL, Sheu K‐FR, Burke JR, Onodera O, Strittmatter WJ, et al. 1997. Polyglutamine domains are substrates of tissue transglutaminase. Does transglutaminase play a role in expanded CAG/poly‐Q neurodegenerative diseases? J Neurochem 69: 431-434. Cooper AJL, Sheu K‐FR, Burke JR, Strittmatter WJ, Gentile V, et al. 1999. Pathogenesis of inclusion bodies in (CAG)n/Qn‐ expansion diseases with special reference to the role of tissue transglutaminase and selective vulnerability. J Neurochem. 72: 889-899. Cooper AJL, Wang J, Pasternack R, Fuchsbauer H‐L, Sheu RK‐F, et al. 2000. Lysine‐rich histone (H1) is a lysyl substrate of tissue transglutaminase: Possible involvement of transglutaminase in the formation of nuclear aggregates in (CAG)n/ Qn expansion diseases. Develop. Neurosci 22: 404-417. Dedeoglu AD, Kubilus JK, Jeitner TM, Matson SA, Bogdanov M, et al. 2002. Therapeutic effects of the transglutaminase inhibitor, cystamine, in a murine model of Huntington’s disease. J Neurosci 22: 8942-8950. De Laurenzi V, MelinoG. 2001. Gene disruption of tissue transglutaminase. Mol Cell Biol 21: 148-155. Dudek SM, Johnson GVW. 1993. Dudek Transglutaminase catalyzes the formation of sodium dodecyl sulfate‐insoluble Alz‐50‐reactive polymers of tau. J Neurochem 61: 1159-1162. Dudek SM, Johnson GVW. 1994. Transglutaminase facilitates the formation of polymers of the b‐amyloid peptide. Brain Res 651: 129-133. Duffel MW, Logan DJ, Ziegler DM. 1987. Cysteamine and cystamine. Methods Enzymol 143: 149-154. Dupuis M, Levy A, Mhaouty‐Kodja S. 2004. Functional coupling of rat myometrial a1‐adrenergic receptors to Gha/ tissue transglutaminase 2 during pregnancy. J Biol Chem 279: 19257-19263.

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Protein Alterations in Mental Retardation

M. A. Junaid . W. T. Brown

1 1.1 1.2 1.3

General Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 Prevalence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 Gender Involvement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261

2 2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.1.5 2.1.6 2.1.7 2.1.8 2.1.9 2.1.10 2.2 2.2.1 2.2.2 2.2.3 2.2.4 2.2.5 2.2.6 2.2.7 2.3 2.3.1 2.4 2.4.1 2.4.2 2.4.3 2.4.4 2.4.5 2.4.6 2.4.7 2.5 2.5.1 2.5.2 2.5.3 2.5.4 2.5.5 2.5.6

Causes of MR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262 Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262 PKU . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262 Creatine Defects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262 Galactosemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 263 Malonyl‐CoA Decarboxylase and Methylmalonyl‐CoA Decarboxylase Defects . . . . . . . . . . . . . . . . 263 HADII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 263 Maple Syrup Urine Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 Hunter Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 Urea Cycle Defects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 Lissencephaly‐1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 Congenital Disorders of Glycosylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 264 Binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265 Fragile X Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265 Renpenning Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 265 Prader–Willi Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 Angelman Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 MASA Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 266 West Syndrome/Partington Syndrome/Aristaless‐Related Homeobox . . . . . . . . . . . . . . . . . . . . . . . . . . 266 a‐Thalassemia/Mental Retardation Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 Allan–Herndon–Dudley Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 Signal Transduction Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 GDP Dissociation Inhibitor 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 X‐Linked Mental Retardation 30 (MRX30/MRX47) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 Rubinstein–Taybi Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 Slit–Robo GTPase‐Activating Protein 3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 Rho Guanine Nucleotide Exchange Factor 6 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 Ras Pathway Mutations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 Holoprosencephaly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 Receptor Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 X‐Linked Mental Retardation 34 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 X‐Linked Mental Retardation 88 (MRX88) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Arginine Vasopressin Receptor 2 (AVPR2) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Insulin Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Hypoparathyroidism–Retardation–Dysmorphism Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Thyroid‐Stimulating Hormone Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

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Protein alterations in mental retardation

2.5.7 2.6 2.6.1 2.6.2 2.7 2.7.1 2.7.2 2.7.3

Thyroid Hormone Receptor b . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Transcription Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Rett Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Mowat–Wilson Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Cytoskeletal Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 Fukuyama Congenital Muscular Dystrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 a7‐Integrin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 3p Deletion Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272

3 3.1 3.2 3.3 3.4 3.5

Newer Approaches to Identify Altered Proteins in MR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 Array Comparative Genomic Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273 Representational Oligonucleotide Microarray Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274 cDNA Array . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274 Oligonucleotide Arrays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276

4

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 277


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General Introduction

Mental retardation (MR) is defined as a neurodevelopmental disability with multiple etiologies (American Psychiatric Association, 2000). To meet the definition, first, the disability should significantly attenuate intellectual functioning and cause reduced cognitive skills. Second, it should have adaptive behavior deficits or impairments in at least two of the following areas: communication, self‐care, home living, social/ interpersonal skills, use of community resources, self‐direction, academic skills, work, leisure, health, and safety. Third, it should have an onset before the age of 18 years. According to this definition, children having an intelligence quotient (IQ) of approximately 70 or below are generally characterized as mentally retarded (Brown, 2002a). The International Classification of Diseases (ICD‐10) characterizes MR as a condition of arrested or incomplete development of the mind, characterized by impairment of skills manifested during the developmental period. ICD‐10 uses all skills that contribute to the overall level of intelligence, i.e., cognitive, language, motor, and social abilities in determining MR (World Health Organization, 2003). MR has traditionally been subclassified depending upon the IQ range into mild (IQ level of 50–55 to approximately 70), moderate (IQ 35–40 to 50–55), severe (IQ 20–25 to 35–40), and profound (IQ less than 20–25). There has been a growing trend to substitute other terms, such as intellectual disability, mental disability, or learning disability, for mental retardation, which implies a static, unchanging condition rather than one that can change over time (Harris, 2006). However, for this chapter, we will use MR as the commonly understood disability.

1.1 Prevalence Epidemiological studies over the years have revealed variable prevalence rates for MR mainly because of the criteria used to define MR. McLaren and Bryson (1987) after a review of literature have reported a combined prevalence of mild and severe MR to be 3–4 per 1000. In United States, different states have a monitoring program that tracks MR along with other developmental disabilities. The most recent Metropolitan Atlanta Developmental Disabilities Surveillance Program (MADDSP), 2006, reported an MR prevalence of 12 per 1000 children aged 8 years (Bhasin et al., 2006). The California Study reported a prevalence of 3.6 per 1000 births for MR cases of unknown causes (Croen et al., 2001). This study however had excluded children with cerebral palsy and autism.

1.2 Gender Involvement The occurrence of MR in either gender depends mainly upon the causative factors. For genetic factors transcribed through the X‐chromosome, there is a clear evidence of higher incidence of MR, in males, because of occurrence of only one X‐chromosome. Thus, even if one copy of an X‐linked gene is defective, the conditions result in MR. It is true that a number of genes residing on the X‐chromosome are involved in MR. However, for other conditions such as Down syndrome and phenylketonuria (PKU) that result from genetic aberration in genes on other chromosomes, the incidences of MR are equally distributed in both genders.

1.3 Diagnosis Historically, MR has been diagnosed based on several assessments of cognitive and behavioral qualities of affected individuals. These assessments are based on interviews with parents, guardians, and caregivers, besides inputs from teachers at schools. Older children are given a standardized test of intelligence (IQ test).

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Commonly used tests include Stanford–Binet Intelligence Scale, Wechsler Intelligence Scales, Wechsler Preschool and Primary Scale of Intelligence, Kaufmann Assessment Battery for Children, Bayley Scales of Infant Development, Woodcock–Johnson Scales of Independent Behavior, and the Vineland Adaptive Behavior Scale (VABS). Diagnosis of MR is made if the child has below average intellectual skills (an IQ below 70–75) and is limited in two or more adaptive skills. More recently, consequent to identification of genetic factors in the etiology of MR, in addition to assessment batteries, the diagnosis is based on maternal blood testing for a‐fetoprotein levels, amniocentesis, and postpartum genetic and enzymatic analyses for a variety of storage diseases.

2

Causes of MR

There are several hundred disorders that possess features of MR. Over 280 genes have so far been identified that are reported to cause MR (Inlow and Restifo, 2004). All the causative factors of MR can be grouped under four categories: prenatal, perinatal, postnatal, and unknown (Kinsbourne and Graf, 2000). Most of the prenatal causes of MR have a genetic basis, with notable exceptions being infections such as rubella, cytomegalovirus infection, or congenitally acquired HIV, alcohol consumption leading to fetal alcohol syndrome, and substance abuse during pregnancy. Perinatal conditions causing MR include hypoxia especially during childbirth, and viral infection by herpes simplex type II. Among the postnatal causes of MR, lead poisoning by far is the most common factor. Earlier, gasoline fuel and paints that contained lead significantly contributed to MR; in recent years, however, there has been decline in these types of poisoning because of measures that eliminated use of lead in gasoline and paints.

2.1 Enzymes 2.1.1 PKU Among the genetic causes, MR is associated with defects in genes that encode practically every type of protein. Genes that encode enzymes are by far the single largest number among various functional categories that results in MR when mutated. So far, over 130 genes have been identified that encode enzymes and whose defective expressions contribute to MR. These include all subclasses of enzymes. A defective enzyme will lead to accumulation of its substrate leading to storage or a key essential metabolite will be lacking that will cause depletion. One of the classical examples is PKU resulting from a defect in metabolism of the amino acid phenylalanine (Folling, 1934). The defective enzyme is phenylalanine hydroxylase (PAH), which is involved in the conversion of Phe to Tyr (Jervis, 1953). Loss of enzyme activity results in elevated levels of Phe especially in the brain, and excretion of phenylketones in urine. PAH is a complex enzyme comprising of the cofactor tetrahydrobiopterin. PKU results not only from the defective PAH but also from defects in the metabolism of the associated cofactor tetrahydrobiopterin. Defects in enzymes involved in the tetrahydrobiopterin cofactor regeneration also cause PKU. Dihydropteridine reductase reduces the transient quinoid dihydrobiopterin generated from tetrahydrobiopterin upon oxidation of Phe to Tyr (Kaufman, 1958). Mutations in the gene encoding the dihydropteridine reductase result in depletion of tetrahydrobiopterin. Unlike in the case of defective PAH, the dihydropteridine reductase defect is not treatable with diet.

2.1.2 Creatine Defects Defects in enzymes involved in the metabolism of creatine are recognized as a new group of rare, inherited disorders that cause MR, seizures, and speech delay. The use of proton magnetic resonance spectroscopy (H‐MRS) resulted in the identification of three inborn errors of metabolism: two creatine biosynthesis errors (arginine/glycine amidinotransferase or AGAT deficiency (MIM 602360) and guanidi-


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noacetate methyltransferase or GAMT deficiency (MIM 601240)) and one creatine transport error (SLC6A8 deficiency (MIM 300036)) (Stockler et al., 1994; Bianchi et al., 2000; Item et al., 2001; Salomons et al., 2001; Cecil et al., 2001). These creatine‐deficiency syndromes (CDS) share the almost‐complete lack of creatine/ phosphocreatine in the brain, as measured by in vivo H‐MRS. Metabolite measurements in urine and plasma indicate the specific type of disorders (e.g., in SLC6A8‐deficient males, the creatine/creatinine ratio is increased in urine, and guanidinoacetate is increased in urine and plasma of GAMT‐deficient patients). The characteristic features in the clinical presentation of all CDS are mental retardation, expressive speech and language delay, and epilepsy (varying from intractable seizures in GAMT‐deficient patients to mild epileptic or febrile seizures in AGAT‐deficient and transporter‐deficient patients). GAMT‐deficient patients and transporter‐deficient patients may show autistic behavior; in GAMT‐deficient patients with a severe phenotype, (extra)pyramidal symptoms are present (Salomons et al., 2003). Female carriers of SLC6A8 mutations have learning disabilities and/or behavioral problems. The genes for the biosynthetic pathway enzymes AGAT and GAMT are inherited in an autosomal recessive fashion, while the gene for the creatine transporter (SLC6A8) is X‐linked. The phenotype results mainly from depletion of creatine in the brain, causing energy imbalance through the creatine phosphate pathway. Clinical symptoms for the two biosynthetic enzyme deficiencies are treatable by supplementation with dietary creatine.

2.1.3 Galactosemia Galactosemia refers to a group of inherited disorders resulting from the inability to metabolize galactose and elevated levels of galactose in the blood that cause damage to the liver, central nervous system, and various other body systems. Mutations in genes encoding enzymes involved in galactose metabolism cause MR. These are inherited as autosomal recessive traits and include galactose‐1‐phosphate uridyl transferase deficiency (Isselbacher et al., 1956) and galactokinase deficiency (Gitzelmann, 1967). A third enzyme deficiency, uridine diphosphate galactose‐4‐epimerase deficiency, was not found associated with MR (Gitzelmann, 1972).

2.1.4 Malonyl‐CoA Decarboxylase and Methylmalonyl‐CoA Decarboxylase Defects Deficiency of malonyl‐CoA decarboxylase, a mitochondrial enzyme, causes mild MR along with vomiting, seizure, hypoglycemia, and metabolic acidosis during a urinary tract infection (Haan et al., 1986; Gao et al., 1999). The urine from patients suffering this disorder contains increased amounts of malonic, methylmalonic, succinic, adipic, glutaric, and suberic acids. The symptoms aggravate when the diet is rich in fat and lysine. Another mitochondrial enzyme, methylmalonyl‐CoA mutase, involved in the isomerization of methylmalonyl‐CoA to succinyl‐CoA causes MR and spastic quadriparesis with dystonia (Shevell et al., 1993).

2.1.5 HADII The multifunctional mitochondrial enzyme 17b‐hydroxysteroid dehydrogenase (17BHSD10) is involved in the metabolism of 2‐methyl‐3‐hydroxybutyryl‐CoA formed from isoleucine and branched‐chain fatty acids (Yang et al., 2005). Besides these, the enzyme also metabolizes both male and female sex‐steroid hormones (He et al., 2000; Ivell et al., 2003). The gene (HADII) encoding 17BHSD10 is localized on the X‐chromosome, and lack of functional activity is associated with progressive neurodegeneration and MR.

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2.1.6 Maple Syrup Urine Disease Maple syrup urine disease (MSUD) results from a deficiency of any of the components of the branched‐ chain a‐keto acid dehydrogenase complex (BCKAD), which is necessary for the breakdown of the branched‐chain neutral amino acids leucine, isoleucine, and valine (Danner et al., 1989; Fisher et al., 1991). Without the BCKAD, these amino acids build up in the body, and if left untreated lead to progressive nervous system degeneration that precipitates MR. There are five genes that encode the BCKAD complex and these are autosomal recessively inherited. The complex comprises of dihydrolipoyl transacylase (E2), branched‐chain a‐keto acid decarboxylase (E1), dihydrolipoamide dehydrogenase (E3), a kinase, and a phosphatase.

2.1.7 Hunter Syndrome Hunter syndrome (HS) or mucopolysaccharidosis type II results from deficient lysosomal iduronate sulfatase activity leading to tissue deposits of mucopolysaccharides and urinary excretion of large amounts of chondroitin sulfate B and heparin sulfate (Danes and Bearn, 1965). The gene encoding iduronate sulfatase is localized on the X chromosome (Wilson et al., 1990).

2.1.8 Urea Cycle Defects Enzymes associated with the urea cycle when defective cause accumulation of ammonia. In these cases, there is severe neurological damage and death results very early in life. Argininosuccinic aciduria is caused by deficiency of argininosuccinase (Westall, 1960; Walker et al., 1990). The gene for argininosuccinase is localized on chromosome 7. Argininosuccinase participates in the urea cycle in ureotelic animals, where it catalyzes the reversible cleavage of argininosuccinic acid into fumarate and arginine. The disease is inherited as an autosomal recessive disorder, manifested by hyperammonemia, which may be life threatening, and accompanied with massive excretion of argininosuccinic acid and its anhydrides. The ornithine transcarbamylase (OTC) deficiency is an X‐linked disorder that causes hyperammonemia leading to brain damage, mental retardation, and death (Michalak and Butterworth, 1997). OTC deficiency is the most common inborn error of urea cycle enzymes in humans. A large percentage of survivors of neonatal OTC deficiency suffer severe developmental disorders, including seizures, MR, and cerebral palsy.

2.1.9 Lissencephaly‐1 The classic lissencephaly‐1 results in developmental delay, myoclonic jerks and spasms, seizures, generalized hypotonia, microcephaly, and dysmorphic facies. The patients show delayed language and motor development and severe mental retardation. The chromosomal aberration was narrowed down to a deletion on chromosome 17, which led to the identification of the gene LIS1 encoding the enzyme platelet‐activating factor acetylhydrolase, PAFAH1B1 (Reiner et al., 1993). The enzyme is involved in the inactivation of platelet‐activating factor by removing the acetyl group from sn‐2 group. Another disorder termed Miller– Dieker lissencephaly syndrome is also caused by mutations in the LIS1 gene (Hattori et al., 1994).

2.1.10 Congenital Disorders of Glycosylation Congenital disorders of glycosylation (CDGs) comprise a group of inherited disorders in which glycosylation of glycoproteins is defective owing to mutations in genes required for the assembly of lipid‐linked oligosaccharides, their transfer to nascent glycoproteins, or in the processing of protein‐bound glycans. The CDGs could be divided into two types, depending on whether they impair lipid‐linked oligosaccharide assembly and


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transfer (CDG type I) or affect trimming of the protein‐bound oligosaccharide or the addition of sugars to it (CDG type II). Among the type I diseases, deficiency of phosphomannomutase‐2 causes a severe phenotype resulting in a multisystem disorder characterized by mental retardation, nonprogressive ataxia, polyneuropathy, hepatopathy during infancy, and growth retardation (van Schaftingen and Jaeken, 1995). Examples of CDG type II comprises of leukocyte adhesion deficiency (LAD) caused by mutations in the gene encoding the GDP‐fucose transporter (Luhn et al., 2001). The affected children have a distinctive syndrome comprising of unusual facial appearance, severe mental retardation, microcephaly, cortical atrophy, seizures, hypotonia, dwarfism, and recurrent infections with neutrophilia. In the absence of the GDP‐fucose transporter protein, several fucosylated proteins and proteoglycans are deficient in these patients. A fucosylated glycoprotein, substance H, that is the precursor of the blood groups A, B, and O is lacking, consequently, these patients manifest the Bombay blood type. The name leukocyte adhesion deficiency is derived from lack of CD15, which is a fucose‐containing cell surface glycoprotein.

2.2 Binding Proteins 2.2.1 Fragile X Syndrome The fragile X syndrome is the most common Mendelian inherited form of mental retardation (Hagerman and Hagerman, 2002). The name fragile X syndrome derives from the presence of a gap or break near the end of the X chromosome at band q27.3, which appears when cells from an affected individual are cultured in a special media. The fragile X syndrome was the first identified example of a rapidly growing list of triplet repeat diseases, including Huntington’s disease, myotonic dystrophy types 1 and 2, Kennedy’s disease, dentatorubral‐ pallidoluysian atrophy, FRAXE syndrome, Friedreich’s ataxia, and spinocerebellar ataxia types 1, 2, 3, 7, 8, 10, 12, and 17. The fragile X syndrome is usually caused by expansion of an unstable CGG repeat region located within the FMR1 (fragile X mental retardation type 1) gene, which inactivates gene expression (Brown, 2002b). This X‐linked form of mental retardation was first recognized as a common and distinct entity in the late 1970s. Current estimates are that approximately 1 in 4,000 males and 1 in 8,000 females in the general population are retarded as a result of the syndrome and that 1 in 300 females is a carrier. Recognizable physical features that are variably present include macroorchidism, large or prominent ears, highly arched palate, narrow midfacial diameter, narrow intereye distance, long face, large head circumference, prominent forehead, facial asymmetry, prominent thumbs, hyper extensible joints, and mitral valve prolapse. Approximately 95% of adult males with fragile X syndrome have an IQ below 70, with an overall mean of 35–50. The FMR1 protein is an RNA‐binding protein and appears to modulate RNA translation of a limited set of mRNAs. Absence of FMRP may lead to impaired synaptic plasticity that may account for the cognitive deficits found in affected individuals. As a result of the trinucleotide expansion, hypermethylation of the CpG repeats ensues that represses the expression of the FMR1 gene. Consequently, the protein FMRP levels are reduced. FMRP is an RNA‐binding protein associated with polyribosomes and may be involved in nuclear export, cytoplasmic transport, and translation of target mRNAs (Bardoni et al., 2001).

2.2.2 Renpenning Syndrome Renpenning syndrome is caused by mutation in the PQBP1 gene that encodes a nuclear polyglutamine‐ binding protein possessing a WW domain (Kalscheuer et al., 2003). Enokido and coworkers (2002) examined the effects of PQBP1 on primary‐cultured cerebellar neurons and concluded that overexpression of PQBP1 inhibits basal transcription in cerebellar neurons and increases their vulnerability to low potassium conditions. Renpenning syndrome is another member of a group that forms part of X‐linked mental retardation. The characteristics include mental retardation, short stature, microcephaly, brachycephaly, spastic diplegia, microgenitalia, and possibly intrauterine growth retardation.

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2.2.3 Prader–Willi Syndrome Prader–Willi syndrome (PWS) is characterized by diminished fetal activity, obesity, muscular hypotonia, mental retardation, short stature, hypogonadotropic hypogonadism, and small hands and feet (Prader et al., 1956). It can be considered to be an autosomal dominant disorder and is caused by deletion or disruption of a gene or several genes on the proximal long arm of the paternal chromosome 15 or maternal uniparental disomy 15, because the gene(s) on the maternal chromosome(s) 15 are virtually inactive through imprinting. PWS is a contiguous gene syndrome resulting from deletion of the paternal copies of the imprinted SNRPN gene encoding the small nuclear ribonucleoprotein polypeptide N (Reed and Leff, 1994), NDN gene encoding the necdin protein (Jay et al., 1997), and possibly other genes.

2.2.4 Angelman Syndrome Angelman syndrome (AS) refers to a condition with features of severe motor and intellectual retardation, ataxia, hypotonia, epilepsy, absence of speech, and unusual facies characterized by a large mandible and open‐mouthed expression revealing the tongue (Angelman, 1965; Bower and Jeavons, 1967). AS is caused because of mutations in the UBE3A gene localized on chromosome 15q11‐q13 that encodes the ubiquitin protein ligase E3A (Kishino et al., 1997; Matsuura et al., 1997). Matsuura and coworkers (1997) identified truncating mutations in patients with AS, indicating that UBE3A is the gene responsible for AS and suggested the possibility of a maternally expressed gene product in addition to the biallelically expressed transcript of the UBE3A gene. Kishino and coworkers (1997) found novel UBE3A mutations in nondeletion/nonuniparental disomy/nonimprinting mutation AS patients. These mutations cause a frameshift and premature termination of translation. These studies suggested that AS is the first recognized example of a genetic disorder of the ubiquitin‐dependent proteolytic pathway in mammals. It also represents an example of a disorder associated with a locus, producing functionally distinct imprinted and biallelically expressed gene products.

2.2.5 MASA Syndrome MASA syndrome refers to clinical features that include mental retardation, aphasia, shuffling gait, and adducted thumbs (Bianchine and Lewis, 1974). The affected gene was identified following demonstration of mutations in the L1 cell adhesion molecule (L1CAM) gene in patients with MASA syndrome by Jouet and coworkers (1994) and by Vits and coworkers (1994). The gene encoding the L1CAM is localized on the X‐chromosome and mutations in L1CAM also causes conditions also known as spastic paraplegia type 1 and X‐linked hydrocephalus (Ruiz et al., 1995).

2.2.6 West Syndrome/Partington Syndrome/Aristaless‐Related Homeobox West syndrome consists of infantile spasms, an electroencephalographic pattern of hypsarrhythmia, and subsequent mental retardation (Feinberg and Leahy, 1977). The gene was mapped to the short arm of the X chromosome (Bruyere et al., 1999) and later identified as ARX (Stromme et al., 2002). ARX is expressed predominantly in the fetal and adult brains, suggesting that ARX protein is important for the maintenance of specific neuronal subtypes in the cerebral cortex and axonal guidance in the floor plate. Two recurrent mutations, present in seven families, resulted in expansion of polyalanine tracts in the ARX protein. These probably caused protein aggregation, similar to other polyalanine and polyglutamine disorders.


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2.2.7 a‐Thalassemia/Mental Retardation Syndrome Weatherall and coworkers (1981) reported the association of hemoglobin H disease and mental retardation in three unrelated patients of northern European descent. Wilkie and coworkers (1990) reported mental retardation and a‐thalassemia owing to deletions of chromosome 16p in eight unrelated patients. The deletion type of the disorder results from omission of approximately 2 Mb from chromosome 16p13.3 region, which involves the hemoglobin a‐1 (HBA1) and a‐2 (HBA2) genes. The nondeletion type of a‐thalassemia/mental retardation syndrome (ATR‐X) is caused by mutations in the ATRX gene (Gibbons et al., 1995). The term X‐linked mental retardation‐hypotonic facies syndrome comprises several syndromes previously reported separately. These include Juberg–Marsidi, Carpenter–Waziri, Holmes– Gang, and Smith–Fineman–Myers syndromes as well as X‐linked mental retardation with spastic paraplegia. All these syndromes are caused by mutation in the ATRX or XH2 gene that encodes helicase 2 and are characterized primarily by severe mental retardation, dysmorphic facies, and a highly skewed X‐inactivation pattern in carrier women (Abidi et al., 2005). The mutations in ATRX cause a generalized dysregulation of gene expression that is responsible for multiple congenital malformations with mental retardation.

2.3 Transporters 2.3.1 Allan–Herndon–Dudley Syndrome Allan and coworkers (1944) described kindred of 24 males affected by severe mental retardation spanning six generations. The patients had hypotonia at birth, but otherwise appeared normal. By 6 months, they developed an inability to hold up the head, leading to the family’s description of the patients as having a ‘‘limber neck.’’ Motor development was markedly reduced, few ever walked, and most had generalized muscular atrophy, joint contractures, and hyporeflexia as adults. Allan–Herndon–Dudley syndrome (AHDS) is caused by mutation in the gene (SLC16A2) encoding the monocarboxylate transporter‐ 8 (Friesema et al., 2004). The transporter protein is involved in the transport of thyroid hormones. In this disorder, serum thyroxin (T4) level is decreased, while serum T3 is increased. Female family members have mild serum thyroid hormone abnormalities but no neurologic manifestations.

2.4 Signal Transduction Proteins 2.4.1 GDP Dissociation Inhibitor 1 GDP dissociation inhibitors (GDIs) are proteins that regulate the GDP–GTP exchange reaction of members of the Rab family, small GTP‐binding proteins of the Ras superfamily, which are involved in vesicular trafficking of molecules between cellular organelles. The Rab proteins undergo activation upon GTP binding, and GTP hydrolysis to GDP inactivates the protein. GDIs slow the rate of dissociation of GDP from Rab proteins and release GDP from membrane‐bound Rabs (Bachner et al., 1995). GDI binds to the small G proteins of the Rab family, particularly the brain‐specific form RAB3A, and modulates their activity and vesicle‐mediated transport. D’Adamo and coworkers (1997) found that GDI is expressed in all parts of the adult brain and, using in situ hybridization analysis, it is detectable in postmitotic neural cells during mouse development, with the same timing and in the same cell types as Rab3a. Unique mutations were found in the RABGDIA gene in affected members of the X‐linked MR 41 and their families. Chelly (1999) referred to the protein mutant in MRX41 as oligophrenin‐2 (OPHN2). Mutations in another X‐linked MR 60 were found to be associated with the gene encoding the oligophrenin‐1,OPHN1 (Billuart et al., 1998). Both OPHN1 and OPHN2 mediate GTP hydrolysis of members of the Rho subfamily, thereby negatively regulating Rho‐GTPase activity. Inactivation of a RhoGAP protein might cause constitutive activation of GTPase targets, which is known to affect cell migration and outgrowth of axons and dendrites in vivo, suggesting an association between cognitive impairment and a defect in a

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signaling pathway that depends on Rho GTPases. Symptoms of MR60 included MR, androgen insensitivity, seizures, ataxia, hypotonia, enlargement of the cerebral ventricles, and cerebellar hypoplasia.

2.4.2 X‐Linked Mental Retardation 30 (MRX30/MRX47) Des Portes et al. (1997) reported a family in which six males in two generations had nonsyndromic X‐linked MR. All affected males had moderate to severe mental retardation without seizures, statural growth deficiencies, or other physical abnormalities. Affected individuals in a multiplex pedigree, previously mapped to Xq22, show a point mutation in the PAK3 gene, which encodes a serine‐threonine kinase, p21‐activated kinase. PAK proteins are crucial effectors linking Rho GTPases to cytoskeletal reorganization and to nuclear signaling. The mutation in PAK3 produces premature termination, disrupting the kinase function. Signal transduction through Rho GTPases and PAK3 may be critical for cognitive function.

2.4.3 Rubinstein–Taybi Syndrome Rubinstein and Taybi (1963) reported a syndrome (Rubinstein–Taybi Syndrome; RSTS) characterized by mental retardation, broad thumbs and toes, facial abnormalities, and which also displays congenital glaucoma. The affected individuals were found to have microdeletions in chromosome 16, and the defective gene identified encodes CREB‐binding protein, CBP (Petrij et al., 1995). CBP is localized in the nucleus and participates as a coactivator in cAMP‐regulated gene expression. It is a potent histone acetyltransferase. When cellular levels of cAMP increase, a cascade of events leads to the induction of genes that contain cis‐ regulatory elements called cAMP‐response elements (CREs). Elevated cAMP levels cause stimulation and nuclear translocation of protein kinase A, which activates the transcription factor CREB (CRE‐binding protein) by phosphorylating it at a single residue, serine‐133 (Gonzalez and Montminy, 1989). The phosphorylated form of CREB binds to CBP (Chrivia et al., 1993). In RSTS patients, these cascades of events are abolished leading to multiple congenital malformations with MR.

2.4.4 Slit–Robo GTPase‐Activating Protein 3 The slit proteins help in neuronal and leukocyte migration through the roundabout transmembrane receptors (ROBO1). Several GTPase‐activating proteins interact with the C terminus of ROBO1. One of these, termed SRGAP3 (for slit–robo GTPase 3), was shown to be deficient in a patient with a balanced de novo translocation t(X;3)(p11.2;p25), hypotonia, and severe mental retardation (Endris et al., 2002). The deficiency of slit‐robo signal transduction pathway prevents correct migration of neurons and their axonal connectivity that are important for normal cognitive function.

2.4.5 Rho Guanine Nucleotide Exchange Factor 6 Rho guanine nucleotide exchange factors (ARHGEFs) belong to a family of cytoplasmic proteins that activate the Ras‐like family of Rho proteins by exchanging bound GDP for GTP. In a large family with nonspecific X‐linked mental retardation (MRX46), mutation in the ARHGEF6 gene was reported (Kutsche et al., 2000). The ARHGEF6 gene encodes a Rho GEF6 protein involved in the Rho GTPase signal transduction, which mediates organization of the cytoskeleton, cell shape, and motility. An actin‐binding protein called PARVB interacts with Rho GEF6 and this interaction is abolished in several MR46 patients (Rosenberger et al., 2003).


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2.4.6 Ras Pathway Mutations Noonan syndrome (NS) is an autosomal dominant dysmorphic disorder characterized by hypertelorism, downward eyeslant, and low‐set posteriorly rotated ears. The disorder is caused in about 50% of cases by mutations in PTPN11, a gene encoding the nonreceptor protein tyrosine phosphatase SHP2, which contains two Src homology‐2 (SH2) domains (Tartaglia et al., 2001). Most of the PTPN11 mutations are in the interacting portions of the amino N‐SH2 and the phosphatase domains, which are involved in switching the protein between inactive and active conformations. Structural analysis of N‐SH2 mutants indicated a significant shift of the equilibrium favoring the active conformation. This finding suggests that gain‐of‐function changes resulting in excessive SHP‐2 activity underlie the pathogenesis of NS. N‐SH2 relays signals from activated receptor complexes to downstream effectors, including RAS. Mutations in KRAS were found in five individuals with Noonan syndrome (Schubbert et al., 2006). Mutations is other members of the RAS/MAPK signaling pathway (Raf, MEK, Hras, and BRAF) have been found in individuals with related disorders associated with developmental delay including Costello and cardio‐ facial‐cutaneous syndromes (Rodriguez‐Viciana et al., 2006; Niihori et al., 2006).

2.4.7 Holoprosencephaly Holoprosencephaly (HPE) is a group of congenital structural forebrain anomalies with craniofacial malformations associated with mental retardation. These disorders are caused by disruption of the sonic hedgehog signaling pathway. One of the causes for HPE is mutation in the gene encoding for the protein sonic hedgehog, SHH, (Roessler et al., 1996). The disorder, HPE3, is caused even by mutations in a single allele as in heterozygous carriers, resulting in insufficient protein levels. SHH is a signal protein that is instrumental in patterning the early embryo. It is expressed in the Hensen node, the floor plate of the neural tube, the early gut endoderm, the posterior of the limb buds, and throughout the notochord. It induces the signal for the patterning of the ventral neural tube, anterior–posterior limb axis, and ventral somites (Riddle et al., 1993; Johnson et al., 1994). Ming and coworkers (2002) have reported another disorder HPE7caused by mutations in patched protein (PTCH). PTCH is the receptor for SHH which normally acts to repress SHH signaling. This repression is relieved when SHH binds to PTCH. Mutations in PTCH decrease SHH signaling, resulting in HPE 7. The mutations could affect the ability of PTCH to bind SHH or perturb the intracellular interactions of PTCH with other proteins involved in SHH signaling. These findings demonstrate genetic heterogeneity associated with the HPE phenotype and show that mutations in different components of a single signaling pathway can result in the same clinical disorder.

2.5 Receptor Proteins 2.5.1 X‐Linked Mental Retardation 34 Carrie and coworkers (1999) demonstrated the importance of interleukin signaling pathways in cognitive function and the normal physiology of the central nervous system. After a thorough investigation of a critical region for X‐linked mental retardation in Xp22.1‐p21.3, they identified a gene expressed in the brain responsible for the nonspecific form of X‐linked mental retardation (MRX34). This gene encodes a 696‐ amino acid protein that has homology to interleukin‐1 receptor accessory proteins (IL1RAP). The results suggested that signal transduction through multifunctional proteins of the immune system may be critical for the development of physiologic processes underlying cognitive function. It was recently shown that the IL1RAP interacts with neuronal calcium sensor‐1 through its specific C‐terminal domain and functions in calcium‐dependent exocytosis.

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2.5.2 X‐Linked Mental Retardation 88 (MRX88) Vervoort and coworkers (2002) described mutations in the angiotensin II‐specific receptor 2 (AGTR2) gene in several patients with MR. These patients completely lack the expression of AGTR2 and displayed MR that ranged from moderate to severe. AGTR2 is abundantly expressed in the fetus and mediates apoptosis through the dephosphorylation of mitogen‐activated protein kinase (Yamada et al., 1996).

2.5.3 Arginine Vasopressin Receptor 2 (AVPR2) A patient with a lifelong history of polyuria, polydipsia, and mental retardation resulting from repeated and prolonged episodes of dehydration was reported, who was shown to have a single‐base pair deletion in the AVPR2 gene, causing a frameshift and premature termination of the protein at codon 270 (Rosenthal et al., 1992). AVPR2 is a G‐protein‐coupled receptor that activates adenylyl cyclase upon binding with G proteins, and maintains the water homeostasis in kidney. Lack of the AVPR2 leads to excessive loss of water, resulting in dehydration, which is responsible for the MR.

2.5.4 Insulin Receptor Two Japanese patients were reported who had nonketotic insulin‐resistant diabetes mellitus with markedly elevated serum insulin values, acanthosis nigricans, hirsutism, and virilism along with mental retardation, short stature, and dental dysplasia. These patients were found to have a defect in the gene encoding the insulin receptor (INSR) that affected the proteolytic maturation of the receptor (Yoshimasa et al. 1988). The INSR is a tetramer comprising of two a‐ and b‐ subunits coded by a single gene and are joined by disulfide bonds. Mutations either in the structural gene or in some of the processing steps lead to insulin resistance that affects the signal transduction cascade and is responsible for the symptoms of MR.

2.5.5 Hypoparathyroidism–Retardation–Dysmorphism Syndrome Congenital hypoparathyroidism in association with growth and mental retardation and seizures has been reported exclusively in children from the Middle East in consanguineous parents (Sanjad et al., 1988). The height, weight, and head circumference scores in affected children are less than two standard deviations from the mean for their ages. These children have identical facies with deep‐set eyes, depressed nasal bridge with beaked nose, long philtrum, thin upper lip, micrognathia, and large, floppy earlobes. Parvari and coworkers (2002) demonstrated that both the hypoparathyroidism–retardation–dysmorphism Syndrome (HRDS) and the autosomal recessive Kenny–Caffey syndrome are caused by deletion and truncation mutations in the TBCE gene, which encodes tubulin‐specific chaperone E. The TBCE gene encodes one of several chaperone proteins required for the proper folding of a‐tubulin subunits and the formation of ab‐tubulin heterodimers. Analysis of diseased fibroblasts and lymphoblastoid cells showed lower microtubule density at the microtubule‐organizing center and perturbed microtubule polarity in diseased cells.

2.5.6 Thyroid‐Stimulating Hormone Receptor A female patient with severe congenital hypothyroidism and absence of circulating thyroglobulin because of TSH unresponsiveness was described (Tonacchera et al., 2000). Serum T4 and T3 concentrations and thyroglobulin were below the sensitivity of the methods, with elevated serum TSH levels. Mutational analysis identified a homozygous mutation in the TSHR gene, resulting in a Thr477Ile substitution in


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the thyroid‐stimulating hormone receptor (TSHR) protein. Immediate family members were heterozygous for the Thr477Ile allele and were unaffected.

2.5.7 Thyroid Hormone Receptor b Phillips et al. (2001) reported a child with extreme thyroid hormone resistance and who presented with goiter, growth retardation, short stature, and deafness. Additionally, the patient had hypotonia, mental retardation, visual impairment, and a history of seizures. Brain MRI showed evidence of demyelination and bilateral ventricular enlargement. The patient had markedly elevated levels of thyroid hormones, T3, T4, and TSH. Molecular analyses of the patient’s DNA identified a single‐base deletion in the TR‐b gene that resulted in a frameshift mutation causing a premature termination codon.

2.6 Transcription Factors 2.6.1 Rett Syndrome Rett (1966) first described a syndrome after observing two girls who exhibited the same unusual behavior. Hagberg et al. (1983) described 35 patients, all girls with a progressive encephalopathy that progressed to severe dementia, autism, loss of use of the hands, and jerky ataxia. Additional neurologic abnormalities intervened insidiously, mainly spastic paraparesis, vasomotor disturbances of the lower limbs, and epilepsy. In sporadic as well as familial patients with Rett syndrome, missense mutations in the MECP2 gene encoding the methyl CpG‐binding protein were found Amir et al. (1999). The mutations were not detected in their obligate carrier mother suggesting that the mother was a germline mosaic for the mutation. The authors suggested that abnormal epigenetic regulation might be a mechanism underlying the pathogenesis of Rett syndrome. Rett syndrome has also been reported for male subjects. Mutations in the MECP2 gene were found in X‐linked MR, referred as MRX16 or MRX79. Orrico et al. (2000) reported a family of a mother and four affected sons, aged 27–40 years, who had normal head size, severe mental retardation with friendly personalities, impaired expressive language development, resting tremors, and slowness of movement. The authors reported a point mutation in the MECP2 gene. Several other groups have also confirmed these findings in other families (Gendrot et al., 1999; Couvert et al., 2001; Yntema et al., 2002; Winnepenninck et al., 2002). Apart from mutations in MECP2, Rett syndrome is also caused by mutations in the CDKL5 gene encoding cyclin‐dependent kinase‐like 5 (Weaving et al., 2004; Tao et al., 2004) and the NTNG1 gene encoding netrin G1 (Borg et al., 2005).

2.6.2 Mowat–Wilson Syndrome Mowat–Wilson syndrome (MWS) is an autosomal dominant complex developmental disorder; individuals with functional null mutations present with mental retardation, delayed motor development, epilepsy, and a wide spectrum of clinically heterogeneous features suggestive of neurocristopathies at the cephalic, cardiac, and vagal levels (Mowat et al., 1998). Mowat‐Wilson syndrome has many clinical features in common with Goldberg‐Shprintzen megacolon syndrome but both disorders are genetically distinct. Wakamatsu et al. (2001) identified nonsense or frameshift mutations in the Smad interacting protein (SMADIP1) or the zinc finger homeobox 1B (ZFXH1B) gene. Expression of wild‐type SMAD1p1 downregulates the transcription of E‐cadherin through binding to both the conserved E2 boxes of the minimal E‐cadherin promoter.

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2.7 Cytoskeletal Proteins 2.7.1 Fukuyama Congenital Muscular Dystrophy In Japan, a number of patients were described who manifest generalized muscle weakness, hypotonia and are unable to walk without support (Fukuyama et al., 1981). All these patients show MR, have seizures, abnormal electroencephalograms and CT scans. In Fukuyama congenital muscular dystrophy (FCMD), the brain malformations include cerebral and cerebellar micropolygyria, fibroglial proliferation of the leptomeninges, hydrocephalus, focal interhemispheric fusion, and hypoplasia of the corticospinal tracts. The majority of these patients show an ancient retrotransposal insertion of tandemly repeated sequences in the FKTN gene, which encodes the fukutin protein. Deficiency of fukutin, a Golgi compartment‐resident protein affects the glycosylation of dystrophin‐associated protein a‐dystroglycan (DAG1), which cannot function properly and elutes from the extracellular surface membrane. DAG1 plays an important role in the assembly of the extracellular matrix in muscle, brain, and peripheral nerves by linking the basal lamina to cytoskeletal proteins (Matsnmura et al., 1993).

2.7.2 a7‐Integrin Mutations in the gene that encodes a7‐integrin were reported in a 4‐year‐old child who developed severe congenital myopathy (Hayashi et al., 1998). This child had severe delayed psychomotor activity along with MR and had severely diminished verbal ability. Expression of the a7‐integrin gene (ITGA7) is developmentally regulated during the formation of skeletal muscle. It is also expressed in the developing nervous system that may be related to psychomotor activity and MR.

2.7.3 3p Deletion Syndrome The gene for the mental retardation in 3p deletion (3p‐) syndrome was identified from an EST with strong homology to the rat Caml1 gene, L1CAM (Wei et al., 1998). The protein encoded is called CALL (cell adhesion L1‐like), which is a novel member of the L1 gene family of neural cell adhesion molecules. The conserved cytoplasmic domain has motifs that are involved in signal transduction pathways. The CALL is expressed regionally in a timely fashion in the central nervous system, spinal cord, and peripheral nervous system during development, lack of which explains the pathology in the disorder.

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Newer Approaches to Identify Altered Proteins in MR

A number of approaches have been successfully applied in the past that resulted in identification of various genes and altered proteins in MR. With the rapid advances made recently in whole genome sequencing, enormous potential exists for the identification of a complete catalogue of MR‐related genes. Development of high‐throughput techniques that evaluate the global expression of the genome will reduce the time needed for identification of gene alterations in MR. These approaches will also help in studying the more complex developmental disorders that have multiple etiologies. The search to resolve the underlying genetic abnormalities in complex developmental disorders, such as autism and schizophrenia, will be assisted by these approaches. Two newer advanced approaches for evaluating defects in gene expression are microarray and proteomic studies. These are finding increasing use in the diagnosis of MR. Both these techniques deal with the global analysis of genome expression. While microarray studies involve genomic DNA or RNA, proteomic studies deal with analysis of global protein expression.


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Genomic DNA copy number alterations are a common feature of many human diseases including mental retardation. Definitive prenatal cytogenetic diagnosis is currently limited to metaphase karyotype analysis of cultured cells obtained by amniocentesis or chorionic villus sampling. Amniotic fluid (AF) samples contain predominantly dying cells. Typically, several weeks are required to promote growth and expansion of the generally low number of viable cells present to allow for metaphase analysis. When particular fetal genetic abnormalities are suspected, additional aberrations—such as deletions, duplications or translocations—can be evaluated using FISH (fluorescence in situ hybridization) analysis with specific DNA probes (Pergament, 2000; Hulten et al., 2003).

3.1 Array Comparative Genomic Hybridization Among the newer molecular techniques, array comparative genomic hybridization (aCGH) allows the detection of microdeletions and microduplications in chromosomes in a high‐throughput fashion. The aCGH is especially well suited for identifying MR‐related alterations, because the underlying abnormality in many MR cases are related to differences in copy number of DNA. The technique is a modification of conventional comparative genomic hybridization (cCGH), which is a molecular cytogenetic technique that detects and maps alterations in copy number of DNA sequences. In the cCGH, the genomic DNA from two samples (patient and normal) is labeled by two different color fluorescent dyes. These DNA samples are hybridized to normal human metaphase chromosomes where sequences from both sources compete for their targets. The images of both fluorescent signals are captured, and ratios of the two hybridization signals are digitally quantified along the entire length of each chromosome. If the chromosomal regions are equally represented in both samples, the color appears as mixture of both fluorescent signals and will have a ratio of 1. Deleted regions will have only one signal and have a ratio below 1, while amplified regions will also have only one signal but of the different color and have a ratio above 1. In this way, a global overview of chromosomal gains and losses throughout the whole sample genome can be obtained. Because cCGH enables analysis of all chromosomes in a single experiment and no dividing cells are required, it has become one of the most popular genome‐scanning techniques. Ghaffari et al. (1998) first showed the potential of the CGH technology in identifying telomeric translocation when they reported chromosomal abnormalities in idiopathic mental retardation in five different patients from three different families. The cCGH suffers from the disadvantage of a reduced ability to detect chromosomal mosaicism, translocations, inversions, and whole‐genome ploidy. Another limitation of the cCGH is the lack of sensitivity, since, the resolution of metaphase chromosomes is of the order of 5–10 Mb. This limitation led to the development of the aCGH. In contrast, in aCGH, the metaphase chromosomes are replaced by cloned DNA fragments of 100–200 kb sizes of known loci. Because of the smaller size, the detection of aberrations is possible in more detail directly onto the genomic sequence. The aCGH is similar to the cCGH in that both use differentially labeled test and reference DNAs. The labeled DNAs are then hybridized to cloned fragments of genomic DNA or complementary DNA (cDNA), which are spotted on a microarray. The DNA copy number aberrations are ascertained by measuring the intensity differences in the hybridization patterns of both DNA. Vissers et al. (2003) applied this approach to identify microdeletions and microduplications, not visible by routine chromosome analysis. In a series of 20 cytogenetically normal patients with mental retardation and dysmorphism suggestive of a chromosomal abnormality, they were able to identify three microdeletions and two microduplications that were previously unidentified. The sensitivity of this approach made possible identifying deletions and duplications as small as 1 Mb.

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3.2 Representational Oligonucleotide Microarray Analysis Sebat et al. (2004) have developed an ultra‐high resolution technique, ROMA (representational oligonucleotide microarray analysis). They have shown that it can define cytogenetic aberrations with extraordinary precision providing an average resolution of 35 kB (Jobanputra et al., 2005). Bianchi et al. (2001) have shown that large amounts of cell‐free fetal DNA is obtainable from the fraction of AF that is primarily discarded. The quality of DNA was comparable to that prepared from cultured AF cells. This DNA predicted correctly the gender of the fetus from the amplification of the male‐ specific FCY gene. In a subsequent article, this group has shown the potential of using cell‐free fetal DNA for rapid prenatal diagnosis of MR using CGH (Larrabee et al., 2004). This group has demonstrated that the cell‐free fetal DNA was suitable for CGH studies to identify whole‐chromosome microduplication and microdeletion in conditions that cause MR, such as trisomy 21 and monosomy X. They also pointed out the advantages of CGH when using cell‐free fetal DNA over normal cytogenetic karyotyping. While cytogenetic karyotyping requires culture of AF cells for several days before significant amounts of cells are present, cell‐ free fetal DNA is obtained immediately at the time of drawing of the AF. A significant proportion of MR cases are caused by microdeletion in the telomeric regions in chromosomes. These regions are characterized by specific sequence units repeated several thousand times. There are a high number of genes that encode proteins in these telomeric regions. Pickard et al. (2004) used two different techniques, subtelomeric fluorescence in situ hybridization (FISH) and multiplex amplifiable probe hybridization (MAPH) to detect MR‐causing abnormalities in a large patient group. A subtelomeric deletion was found on chromosome 4q35.2, which also shows good linkage evidence (LOD score of 3.2 for microsatellite marker D4S1652) from an extended Australian kindred affected with bipolar affective disorder and mild MR. The deletion spans about 3 Mb and contains at least ten identifiable transcripts. Further gene association studies may be required to identify the candidate psychiatric illness gene at 4q35. Yobb et al. (2005) have utilized four different techniques to identify microduplication in patients with 22q11.2 deletion syndrome. These included FISH, microsatellite markers, real‐time PCR, and MAPH. The group was able to identify microduplications of chromosome 22q11.2 that may contribute to additional diversity in the 22q11.2 deletion syndrome. Cri du chat syndrome occurs owing to deletion involving chromosome 5p. In a study of 94 patients carefully evaluated for the presence of the characteristic cry, speech delay, facial dysmorphology, and level of MR, aCGH has narrowed down the MR region into roughly three areas (Zhang et al., 2005). Deletions of MRI, a 1.2‐Mb region overlapping the previously defined cri du chat critical region but not including MRII and MRIII, produced a moderate level of retardation. Deletions restricted to MRII, located just proximal to MRI, produced a milder level of retardation, whereas deletions restricted to the still‐more proximal MRIII produced no discernible phenotype. However, MR increased as deletions that included MRI extended progressively into MRII and MRIII, and MR became profound when all three regions were deleted. Like other techniques, aCGH studies of the human genome often reveal false negative and false positive results because of using large insert clones as probes. This has raised important concerns regarding the suitability of this approach for clinical diagnostic applications. To circumvent this problem, a computational method, Smith–Waterman algorithm previously developed to identify genomic regions with unusual properties, was used that makes no distributional assumptions about the data to identify putative copy‐ number changes and determine their statistical significance (Price et al., 2005). This method of data analysis represents a way to overcome the problems of low sensitivity and specificity associated with aCGH. The method is adoptable for data from any type of array probe and for all regions of the genome provided there is accurate positional information across a contiguous section of chromosome available.

3.3 cDNA Array The cDNA array methodology is a powerful tool for the simultaneous analysis of mRNA expression levels of a large number of genes. The use of this technology in human diseases characterized by alterations in the genome structure is finding increasing use. A DNA microarray consists of an orderly arrangement of DNA


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fragments representing the genes of an organism. Each DNA fragment represents a gene, assigned a specific location on the array, usually a glass slide, and then microscopically spotted to that location. Over 30,000 spots can be placed on one slide using robotic spotters, allowing analysis of virtually every gene present in a genome. The main advantage of microarrays is that the spots are single stranded DNA fragments strongly attached to the slide, allowing cellular DNA or RNA to be fluorescently labeled and hybridized to the complementary sequence on the array. By exposing the microarray to a fluorescently labeled sample the DNA or RNA that hybridizes will be identifiable as glowing spots on the array, while the spots that have nothing hybridized will not be visible. Two main types of microarrays, cDNA arrays and oligonucleotide arrays, offer different approaches of microarray technology. In cDNA arrays the probes are larger pieces of DNA that are complementary to the genes of interest. These cDNA probes can be generated from a commercially available cDNA library ensuring a close representation of the entire genome of an organism on the array. Alternatively, specific primers can be used to amplify specific genes from genomic DNA to generate the cDNA probes. A separate PCR reaction must be performed for each gene, although these reactions can be done in parallel. The cDNA probes prepared are then mechanically spotted onto a glass slide. Using cDNA microarray with human primary cell cultures of T lymphocytes prepared from normal and Down syndrome subjects, Giannone et al. (2004) have shown over expression of the superoxide dismutase (SOD1), MHC DR b3 (HLA‐DRB3), GABA receptor Ag2 (GABRG2), acetyltransferase coenzyme A2 (ACAT2), and ras suppressor protein 1 (RSU1) genes. Clustering of overexpressed genes in patients and controls showed that the expression is restricted to chromosome 21 genes, which reiterated the specific gene dosage theory in the pathogenesis of the DS phenotype.

3.4 Oligonucleotide Arrays Oligonucleotide arrays use small 25 base pair gene fragments as the DNA to be spotted onto an array. In all, 11 to 16 copies of this DNA are spotted for each gene to be placed on the array. These DNA probes are selected to have little cross‐reactivity with other genes so that nonspecific hybridization will be minimized. To prevent nonspecific hybridization, a second probe that is identical to the first except for a mismatched base at its centre is placed next to the first. This is called the Perfect Match/Mismatch (PM/MM) probe strategy. Any background hybridization with the MM probe is subtracted from the PM probe signal, which results in perfect hybridization. Mao et al. (2005) have used oligonucleotide arrays with over 20,000 transcripts to compare gene expression in Down syndrome fetal tissues versus controls. These investigators have shown that in Down syndrome, there is a primary transcriptional effect of disruption of chromosome 21 gene expression, without a pervasive secondary effect on the remaining transcriptome. There were tissue‐ and cell‐specific changes of gene expression in trisomy 21 during fetal development. The data revealed changes in few of the transcripts derived from nonchromosome 21 genes; however, large‐scale disruption of the genome at other chromosomes was absent. In another approach, microarray of gene expression analysis was studied using the Affymetrix Mu11K high‐density oligonucleotide array, followed by validation of the data using quantitative RT‐PCR (QPCR) in a mouse mode for lysosomal storage disease (Brooks et al., 2002). Improvement and reversal of cognitive and behavioral functions were tested using viral‐based vectors encoding the defective genes. Several genes involved in memory and learning were found upregulated upon viral transfer of defective genes. These included PIPTN, encoding the a isoform of phosphatidylinositol transfer protein (PITPa), C/EBPd, increasing as a result of stimulation of cAMP signaling in hippocampal neurons, EGR2, important for peripheral nerve myelination, CYR61, encoding an extracellular matrix protein found in the human CNS, and FE65, whose product interacts with the g‐secretase‐liberated tail of amyloid precursor protein (APP) and with the histone acetyltransferase Tip60 (41), which are necessary for axonal migration. Although, oligonucleotide arrays were primarily useful for SNP analysis, these are also adaptable for assessing the DNA copy number, large‐scale linkage analysis, and whole‐genome association studies. As

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such, they offer great potential for use in identifying unknown genes in MR. The fluorescent signal from the oligonucleotide array varies in proportion to both decreases and increases in copy number.

3.5 Proteomics Proteomics is a powerful tool in the postgenomic era for evaluating simultaneous global protein expression. The technique requires combination of different methodologies to study the expression profiles of all the cellular proteins at a given time. Proteomics relies on separation of total proteins preferably by two‐ dimensional (2D) gel electrophoresis using immobilized pH gradients. The separated proteins are stained by any of a number of sensitive staining procedures, including silver and fluorescent staining. The images are compared between normal and diseased (or test) samples, and abnormal proteins following enzymatic digestion are identified either by matrix‐assisted laser desorption/ionization time‐of‐flight mass spectrometry (MALDI‐TOF‐MS) or by liquid chromatography followed by tandem MS mass spectrometry/mass spectrometry (Langen et al., 1999). Several other modifications of proteomics are available for tackling specific issues relating to different sets of proteins. Autism is a highly heterogeneous neurodevelopmental disability with a strong indication of genetic factors in the etiology. An autism concordance rate of approximately 90% in monozygotic twins versus 10% in dizygotic twins indicates it has a high degree of heritability (Bailey et al., 1995). Although highly heritable, it is a genetically complex disorder characterized by deficits in language and in reciprocal social interactions, combined with repetitive and stereotypic behaviors. A number of whole‐genome wide genome scans in many laboratories have revealed inconsistent results, although some degree of reproducibility has recently been reported for loci on chromosomes 7 and 17 (Cantor et al., 2005). The evidence of multigenic components, with perhaps 10–15 contributing genes in the etiology of autism (Risch et al., 1999), makes it difficult to use conventional positional cloning approaches. We have successfully utilized proteomics to identify aberrant proteins in autopsied brains of patients with autism. Using autopsy brain tissues from autism and control subjects, we have shown that an altered form of glyoxalase I (Glo1) determines the predisposition to autism. Subsequently, the protein aberration was shown to be the result of a SNP in the coding region that causes Ala111Glu change in the protein, which is responsible for reduced enzyme activity (Junaid et al., 2004). MS is not only applicable to larger biomolecules such as proteins but is also applicable to small metabolites in biological pathways. Wu et al. (2004) have shown that tandem MS in a metabolomics‐ guided screening can identify metabolite differences in N‐ethyl‐N‐nitrosourea‐treated (ENU‐treated) mice. These mice upon ENU treatment develop marked elevation of blood branched‐chain amino acids (BCAAs), ketoaciduria, and clinical features resembling human maple syrup urine disease, caused by the deficiency of branched‐chain a‐keto acid dehydrogenase (BCKD) complex. Another advantage of proteomics is the ability to perform quantitative analyses of the expression of proteins. Using a MECP2‐knockout mouse model for Rett syndrome, Matarazzo and Ronnett (2004) have demonstrated regional variation in protein expression in olfactory lobes. The MECP2 gene encodes MeCP2 protein, which is a transcriptional repressor that binds to methylated cytosine residues in the CpG islands. Defective MeCP2 is thus expected to affect a wide spectrum of protein translations because of defective transcription. As expected, these investigators have found widespread defects in protein expression owing to differences in mRNA levels and posttranslational modifications in the knockout mice. All of the proteins identified were not part of a single functional family of proteins and one chromosomal location. The identified proteins could be regrouped into five main groups, corresponding to cytoskeleton arrangement (CRMP2, CAPG, lamin C), chromatin modeling (histones H3, H2B, and 1H2AK), energy metabolism (cytochrome b5, isovaleryl‐CoA dehydrogenase, creatine kinase), cell signaling (calretinin, 14‐3‐3 b and z), and neuroprotective (GRP78, GSTO1) functions.


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Conclusions

The existence of the concept of MR in humans dates back several thousand years. The presence of a separate holding facility for such individuals was known in 1247 in London (Scheerenberger, 1983). During the last century, enormous strides have been made in the MR field, with respect to the identification of the various causative factors responsible, treatment strategies, and ultimately in the prevention of the disability. To date, over 280 genes that influence MR in one or other aspects have been successfully identified using diverse technologies. There may be many more MR‐causing genes identified in the near future, a task that will consume less time and effort than has taken so far owing to the availability of the completed human genome and high‐throughput DNA copy number and aberration analyses as well as RNA and protein expression technologies. Traditionally, the molecular mechanisms underlying defects in the single gene were easier to identify and understand. A more challenging area is in unraveling the causative factors and associated pathways in complex psychiatric developmental disorders such as autism, schizophrenia, depression, and attention deficit hyperactive disorder. The complex developmental disorders are hypothesized to result from disruption in interactions of multiple genes. Genes that contribute toward major phenotypic characteristics may be more easily identified than those that only marginally affect minor traits. Conventional methods such as association studies to narrow chromosomal loci by linkage analyses may be formidable in delineating defects in complex disorders. In these instances, high‐throughput functional genomic approaches including, but not limited to, microarray and proteomics will eventually prove successful. SNP and cDNA microarrays will refine the effective coverage of whole‐genome scans for the detection of chromosomal aberrations. Currently, SNP array analyses can simultaneously study half a million SNPs that are spaced out a few hundred bases apart in the entire genome. Similarly, oligonucleotide arrays are available that cover over 50,000 transcripts. Both these approaches will extend the scope of whole‐genome scans for identifying chromosomal abnormalities. Complementary to these approaches is the study of global analysis of protein expression by the use of proteomics. Proteomics will have the advantage of increased stability of proteins in tissues in comparison to nucleic acid‐based technologies that rely on the levels of DNA and RNA that are more rapidly degraded. Once a defect in a gene or protein is identified to be responsible for MR, the subsequent elucidation of the molecular mechanism could become a formidable task. In certain cases, such as Fragile X syndrome and Huntington disease, while the causative factors have been known for some time, the functional characterization of the proteins still remains obscure. Functional characterization of the proteins is necessary ultimately to devise effective therapeutic interventions for MR conditions.

Acknowledgments Funding for this research was provided in part by NYS OMRDD and by NIH grant NS40691.

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9

Protein Sulfation

S. Hemmerich

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 284

2

Sulfation Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 286

3

The Human Sulfotransferase Complement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287

4

Tyrosine Sulfation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291

5 5.1 5.2 5.3 5.4 5.5

Sulfation of O‐linked Glycans in Leukocyte Adhesion and Inflammation . . . . . . . . . . . . . . . . . . . . . 291 Sulfated Glycans in Lymphocyte Recirculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291 Sulfated Glycans in Chronic Inflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291 Endothelial Sulfotransferases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 292 Induction of Endothelial Sulfotransferases in Chronic Inflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293 Other Immunologically Important Sulfotopes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293

6 6.1 6.2 6.3

Heparan Sulfate Proteoglycans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 294 Composition and Biosynthesis of Heparan Sulfate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 294 Biological Roles of HS‐Associated Sulfotopes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 294 Endosulfatases Involved in HS Remodeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 294

7

Sulfation in Regulation of Hormone Half‐Life . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 295

8 8.1 8.2 8.3 8.4

Sulfation in the Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296 Sulfated Glycolipids in Myelin Structure and Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296 Chondroitin Sulfate in CNS Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 297 Heparan Sulfate Proteoglycans in Axon Guidance and Targeting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298

9

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298

#


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Protein sulfation

Abstract: Sulfation of glycoproteins, glycolipis, and proteoglycans is a post‐translational modification fundamental to developmental biology, immunology, and neurobiology, as well as disease processes in cancer, inflammation and central nervous system disorders. The enzymes at the heart of biological sulfation are the Golgi compartment‐associated sulfotransferases which decorate their substrates with specific sulfated epitopes (sulfotopes) that eventually become exposed to the extracellular environment and comprise novel binding sites for biological effector molecules facilitating cell‐cell and cell‐matrix communication. This review discusses a number of well defined sulfotopes with important biological roles, such as found on the ligands for the proinflammatory adhesion molecules L‐ and P‐selectin, on pituitary gland hormones, on chemokine receptors involved in inflammation and virology, as well as on proteoglycans involved in hemostasis, growth factor mobilization and neural plasticity. Regulation of sulfotopes occurs often at the level of the underlying sulfotransferase(s), and in some cases also by specific endo‐sulfatases. This review also classifies the human sulfotransferase complement and discusses the potential of sulfotransferase inhibition for novel therapies of important human diseases. List of Abbreviations: APS, adenosine 50 ‐phosphosulfate; CAM, chorioallantoic membrane; CIA, collagen‐ induced arthritis; CSPGs, chondroitin sulfate proteoglycans; CST, cerebroside ST; EAE, experimental autoimmune encephalitis; FGFs, fibroblast growth factors; GAGs, Glucosaminoglycans; GlcNAc, N‐acetylglucosamine; GalNAc4S, GalNAc‐4‐O‐sulfate; GlcNAc6S, GlcNAc 6‐O‐sulfate; GST, Gal/GalNAc/GlcNAc‐ 6‐O‐ST; HEV, high endothelial venule; HS, heparan sulfate; HSPGs, heparan sulfate proteoglycans; IdoA, Iduronate; LH, Lutropin; MS, multiple sclerosis; PAPS, 30 ‐phosphoadenosine 50 ‐phosphosulfate; PDGF, platelet‐derived growth factor; PNAd, peripheral vascular addressin; PSGL‐1, P‐selectin glycoprotein ligand 1; RA, rheumatoid arthritis; sLex, sialyl Lewis x; STs, Sulfotransferases; TSH, Thyrotropin; VEGF, vascular endothelial growth factor

1

Introduction

Following the sequencing of the human genome in 2001 (IHGSC, 2001; Venter et al., 2001) and ongoing refining and interpretation of the data (IHGSC, 2004), it has become clear that in genetic terms a human with 20,000–25,000 genes is about as complex as the tiny roundworm Caenorhabditis elegans (19,500 genes) and significantly less complex than some common crops such as rice (45,000 genes) and maize (50,000 genes). In order to resolve this apparent conundrum, Venter and colleagues (Venter et al., 2001) proposed, already in 2001, that the far lower number of human genes than previously predicted might be compensated for by combinatorial diversity generated at the level of posttranslational modification. Phosphorylation and glycosylation are the most widely recognized posttranslational modifications in biology; however, sulfation is another posttranslational modification whose important biological roles have become appreciated over the last decade. Sulfation involves transfer of a sulfate moiety from a high‐energy nucleotide donor known as 30 ‐phosphoadenosine 50 ‐phosphosulfate (PAPS) to a nucleophilic hydroxyl or amino group on metabolites, tyrosine residues in proteins, and glycans in glycoproteins, glycolipids, or proteoglycans. Mechanistically, sulfation is homologous to phosphorylation (> Figure 9‐1) and involves similar transfer mechanisms (Kakuta et al., 1998b). Sulfotransferases (STs) are the enzymes that transfer sulfate from PAPS to the sulfate acceptor, analogous to the kinases (phosphotransferases) in phosphorylation reactions. Sulfatases are enzymes that hydrolyze sulfate from sulfated molecules, analogous to but mechanistically distinct from the phosphatases (Lukatela et al., 1998; Roeser et al., 2006), their counterparts in dephosphorylation processes. In vertebrates, STs occur as cytosolic enzymes and membrane‐bound enzymes associated with the Golgi compartment. The cytosolic enzymes play important roles in detoxification processes as well as hormone and neurotransmitter metabolism; they have been reviewed extensively elsewhere and are not discussed here (Falany, 1997; Kauffman, 2004; Gamage et al., 2006). In contrast, the Golgi compartment‐ associated STs transfer sulfate onto nascent glycoprotein, proteoglycan, and glycolipid substrates, while these are being shuttled through the Golgi compartment on their way toward expression on the plasma membrane or secretion into the extracellular space. The specific sulfation modifications implemented by the Golgi compartment enzymes onto their substrates, also termed sulfotopes, frequently constitute novel

Protein sulfation

9

. Figure 9‐1 Analogy between sulfation and phosphorylation

binding sites for adhesion molecules, growth and differentiation factors, chemokines, and other biomolecules fundamental to cell–cell and cell–matrix communications. In many cases, specifically on glycosaminoglycan substrates, many structurally and functionally distinct, sometimes also overlapping, sulfotopes are generated on the same substrate (> Figure 9‐2). These sulfotopes and their inherent biological effector functions can further be modulated by another class of enzymes known as sulfatases, which remove specific

. Figure 9‐2 Sulfotopes on glycoproteins and proteoglycans

285

286

9

Protein sulfation

sulfate residues from their substrates. Over the last 15 years, protein sulfation has become recognized as fundamental to many physiological and pathological processes in immunology, endocrinology, developmental biology, and neurobiology. As protein sulfation most commonly occurs on carbohydrate residues within glycoconjugates (glycoproteins, glycolipids, and proteoglycans), its study is linked intrinsically to the study of structure and function of the underlying carbohydrate structures, within a rapidly evolving branch of the life sciences known as glycobiology (Varki and Chrispeels, 1999).

2

Sulfation Pathways

Although sulfated glycol‐conjugates exert their biological effector functions on the cell surface or within the extracellular space, biological sulfation (> Figure 9‐3) commences with uptake of inorganic sulfate from the extracellular medium through plasma membrane‐embedded sulfate channels or sulfate transporters (Markovich, 2001). Some of these sulfate channels show a remarkably restricted expression pattern and may regulate cell‐type specific sulfate metabolism (Girard et al., 1999), rendering them attractive targets for possible pharmacological intervention. Once inside the cytosol, inorganic sulfate is conjugated to ATP through the catalytic action of ATP sulfurylase, resulting in an intermediate adenosine 50 ‐phosphosulfate (APS), which is immediately further phosphorylated at the 30 position of its ribose to yield 30 ‐phosphoadenosine 50 ‐phosphosulfate (PAPS). PAPS is the universal sulfate donor for all sulfation processes in biology. Formation of APS is an energy‐intensive process that requires coupling to an energy‐yielding reaction. In bacteria, ATP sulfurylase (cysD) and APS kinase (cysC) reside in two different components of a ternary complex, which also contains a GDPase (cysN) (Mougous et al., 2006). The latter provides the energy required to drive ATP sulfurylation. In vertebrates, ATP sulfurylation and APS phosphorylation are

. Figure 9‐3 Sulfation pathways and generation and modulation of sulfotopes in vertebrate cells

Protein sulfation

9

catalyzed by two isoforms of a single chain enzyme known as PAPS synthase (Venkatachalam, 2003). However, the mammalian enzymes are not coupled to GTP hydrolysis, rather this reaction is driven in the physiologic direction by the hydrolysis of pyrophosphate catalyzed by an ubiquitous inorganic pyrophosphatase (Segal et al., 1987). In animal cells, PAPS serves exclusively as a sulfate donor for STenzymes, while in bacteria, fungi, and plants, PAPS can be reduced to essential sulfur metabolites such as cysteine and methionine through a process known as sulfate assimilation (Bick and Leustek, 1998; Kopriva, 2006). In the mammalian cytosol, PAPS can be utilized directly by cytosolic STs, which are involved in detoxification processes (through sulfation of toxins for increased solubility and secretion into urine) as well as in the metabolism of steroids and catecholamines. More importantly for the purpose of this review, PAPS can be translocated to the Golgi compartment lumen through a recently identified Golgi compartment membrane‐embedded PAPS transporter orthologous to the Drosophila melanogaster slalom protein (Kamiyama et al., 2003). In the Golgi compartment, membrane‐embedded STs then transfer sulfate from the Golgi compartment‐resident PAPS to their glycoprotein, proteoglycan, or glycolipid substrates as they are being processed through the Golgi compartment on their way to the plasma membrane or toward being secreted. Because of the particular arrangement of the STs and their substrates within the Golgi compartment (the ST catalytic domains always point to the Golgi compartment lumen), the sulfated aspect of the product, operationally termed sulfotope, will eventually always face the extracellular space, whether associated with the plasma membrane or on a secreted product. As mentioned above, the resulting sulfotopes mediate the downstream biological effects of glycoprotein (and glycolipid) sulfation in cell–cell and cell–matrix communications. Sulfated glycoconjugates, specifically glycosaminoglycans, are degraded through lysosomal pathways involving lysosomal exosulfatases, the absence of which can lead to a number of serious genetic disorders collectively known as lysosomal storage diseases (Freeman and Hopwood, 1992). However, at least one specific sulfotope found on heparin‐sulfate‐type proteoglycans can be directly modulated by a novel type of extracellular endosulfatases, which can remove a critical sulfate moiety from this sulfotope and thus mobilize previously matrix‐bound growth and differentiation factors (discussed in detail in > Sect. 5). The different players in biological sulfation offer a number of target opportunities for pharmacological intervention into pathological processes involving sulfation: Sulfotopes are attractive targets for therapeutic antibodies as they are exposed to the extracellular space. Sulfotopes are less attractive as targets for small‐molecule drugs because of the general hydrophilic nature of sulfated glycans. Golgi compartment‐ associated STs are intracellular enzymes, and therefore only accessible to cell‐permeant small‐molecule drugs. As STs feature structural and mechanistic similarities to kinases (Kakuta et al., 1998, 1999), lessons learned from kinase inhibitor discovery may also apply to this class of enzymes. Extracellular sulfatases in turn can be targeted with small molecules as well as antibodies, and sulfate channels, though poorly validated to date, also pose attractive targets for small‐molecule drugs or biologics. Examples of such target opportunities are discussed below.

3

The Human Sulfotransferase Complement

Biological sulfation was first discovered by Baumann in 1876 in sulfated metabolites in urine (Baumann, 1876). Karl Meyer and colleagues described sulfated glycoconjugates (heparin, chondroitin sulfate, and dermatan sulfate) in the 1930s. Tyrosine sulfation was discovered in 1954 in fibrinogen (Bettelheim, 1954) and in the 1960s enzyme preparations catalyzing sulfation of proteoglycans were first reported. The first cDNA encoding a Golgi compartment‐associated ST, heparin deacetylase N‐ST 1, was cloned in 1992 (Hashimoto et al., 1992). STs share a conserved amino acid sequence motif that binds the universal sulfate donor PAPS (Kakuta et al., 1998a&b; Fukuda et al., 2001). The presence of this conserved motif has been used to identify genes encoding novel carbohydrate STs (Fukuda et al., 2001). Through the availability of the human genome sequence, most, if not all, of the human ST genes have been identified, and many have now been expressed as recombinant proteins and characterized. There are at least 50 genes known to encode, with splice variants, a somewhat larger number of enzymes (> Table 9‐1). These enzymes can be organized into families characterized by sequence homology and functional properties (> Figure 9‐3). One of these families comprises the cytosolic STs, referred to in the introduction, that catalyze sulfation

287

Gal/GalNAc/GlcNAc 6STs

TPSTs

Cytosolic sulfotransferase 1D pseudogene Estrogen sulfotransferase Dehydroepiandrosterone sulfotransferase 3b‐hydroxysteroid sulfotransferase Cytosolic sulfotransferase 4A Cytosolic sulfotransferase 6B Protein tyrosine sulfotransferase‐1 Protein tyrosine sulfotransferase‐2 Chondroitin 6‐O‐sulfotransferase 1 Keratan sulfate galactose 6‐O‐ sulfotransferase N‐acetylglucosamine 6‐O‐ sulfotransferase L‐selectin ligand sulfotransferase Intestinal GlcNAc 6‐O‐sulfotransferase Corneal GlcNAc 6‐O‐sulfotransferase Chondroitin 6‐O‐sulfotransferase 2 NCAG1 similar to sulfotransferase

Catecholamine sulfotransferase Thyroid hormone sulfotransferase Cytosolic sulfotransferase family 1C

Enzyme Phenol sulfotransferase

Accession # NP_001046 NP_001045 NP_003157 NP_055280 NP_001047 NP_006579 DAA01771 NG_002642 NP_005411 NP_003158 NP_004596 NP_055166 DAA01772 NP_003587 NP_003586 NP_004264 NP_003645 NP_004258 NP_005760 NP_036258 NP_067628 NP_063939 NP_115536

Gene SULT1A1 SULT1A2 SULT1A3 SULT1B1 SULT1C1 SULT1C2 SULT1C3 SULT1DP SULT1E1 SULT2A1 SULT2B1 SULT4A1 SULT6B1 TPST‐1 TPST‐2 GST‐0 GST‐1 GST‐2 GST‐3 GST‐4a GST‐4b GST‐5 GST‐X

16q22.2 16q22.1 16q22.1 Xp11.3 18q22.1

3q24

19q13.33 22q13.31 2p22.2 7q11.21 22q12.1 10q22.1 11p11.2

4q13.3 19q13.33

Chromosome locus 16p11.2 16p11.2 16p12.2 4q13.3 2q12.3 2q12.3 2q12.3 4q13.3

41–386 40–390 39–395 100–486 861–1222

163–530

Entire peptide sequence Entire peptide sequence Entire peptide sequence 62–377 66–356 131–479 59–411

Entire peptide sequence Entire peptide sequence

ST‐domain used for alignment to generate 9‐4 Entire peptide sequence Entire peptide sequence Entire peptide sequence Entire peptide sequence Entire peptide sequence Entire peptide sequence Entire peptide sequence Entire peptide sequence > Figure

9

Family Cytosolic sulfotransferases

. Table 9‐1 The sulfotransferase proteome in humans

288 Protein sulfation

Heparan deacetylase N‐ sulfotransferase

Galactosylceramide sulfotransferase Glycoprotein b‐Gal 3‐sulfotransferase b‐Galactose‐3‐O‐sulfotransferase 3 Galb1!3GalNAc 30 ‐sulfotransferase HNK‐1 sulfotransferase Chondroitin‐4‐sulfotransferase

NDSTs

Gal3STs

GalNAc4STs

GalNAc 4‐sulfate 6‐O‐sulfotransferase

GalNAc 4‐O‐sulfotransferase

Heparan sulfate 6‐O‐sulfotransferase

6OSTs

3OSTs

Dermatan sulfate 2‐O‐sulfotransferase Heparan sulfate 2‐O‐sulfotransferase Heparan sulfate 3‐O‐sulfotransferase

2OSTs

DS‐2OST HS‐2OST HS‐3OST‐1 HS‐3OST‐2 HS‐3OST‐3A HS‐3OST‐3B HS‐3OST‐4 HS‐3OST‐5 HS‐3OST‐6 HS‐6OST‐1 HS‐6OST‐2S HS‐6OST‐2L HS‐6OST‐3 NDST‐1 NDST‐2 NDST‐3 NDST‐4 Gal3ST‐1 Gal3ST‐2 Gal3ST‐3 Gal3ST‐4 HNK1ST C4ST‐1 C4ST‐2 C4ST‐3 GalNAcST1 GalNACST2 GalNAc4S6ST

NP_005706 NP_036394 NP_005105 NP_006034 NP_006033 NP_006032 XP_056254 AAN37737 AAK61299 NP_004798 NP_671704 NP_671703 NP_703157 NP_001534 NP_003626 NP_004775 NP_072091 NP_004852 NP_071417 NP_149025 NP_078913 NP_004845 NP_060883 NP_061111 NP_690849 NP_071912 NP_113610 NP_055678 13q32.1 5q33.1 10q22.2 4q26 4q26 22q12.2 2q37.3 11q13.2 7q22.1 2q11.2 12q23.3 7p22.3 3q21.3 19q13.11 18q11.2 10q26.13

6q25.1 1p22.3 4p15.33 16p12.2 17p12 17p12 16p12.1 6q21 16p13.3 2q21.1 Xq26.2

105–406 49–307 110–367 148–406 133–399 208–471 86–346 86–346 55–311 79–410 220–605 73–459 139–471 599–882 598–884 590–873 589–872 72–423 48–398 59–431 63–486 79–256 76–352 119–414 61–341 151–424 168–438 251–561

Protein sulfation

9 289

290

9

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of small metabolites and are not discussed here. All other STs are Golgi compartment‐associated membrane‐bound enzymes catalyzing sulfation of glycoproteins, glycolipids, and proteoglycans. Two of theses enzymes, TPST‐1 and TPST‐2, catalyze sulfation of tyrosine on glycoproteins including adhesion molecules and chemokine receptors, the remainder catalyze regioselective sulfate additions to specific carbohydrate moieties in glycoconjugates. This latter class of glycosyl‐STs boasts the greatest degree of structural and functional diversity within the human ST complement, also termed operationally the human sultome (> Figure 9‐4). They create a large variety of specific sulfotopes that play important roles in cell adhesion and migration, cell proliferation and differentiation, regulation of hormone half‐life, and specifically in neurobiological processes.

. Figure 9‐4 The human sulfotransferase (ST) complement (human sultome)—The abbreviations for the different enzymes and families refer to > Table 9‐1. Protein sequences encoded by the known human ST genes were aligned with the CLUSTALX algorithm (Thompson et al., 1997). The alignment of Golgi compartment STs was done using only the predicted catalytic domains (Hemmerich et al., 2004). The TreeView program (Page, 2005) was used to generate the depicted phylogenetic tree. The scale bar corresponds to 0.1 change per nucleotide

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Tyrosine Sulfation

As mentioned above, tyrosine sulfation was first discovered in fibrinogen (Bettelheim, 1954), where it may enhance binding to thrombin exosites (Meh et al., 2001). More importantly, tyrosine sulfation occurs on a leukocyte adhesion receptor known as P‐selectin glycoprotein ligand 1 (PSGL‐1) as well as on the extracellular aspect of a number of chemokine receptors including CXCR‐3, CXCR‐4, CCR‐2, and CCR‐5 (Kehoe and Bertozzi, 2000). In the case of PSGL‐1, where three tyrosine residues are sulfated, sulfation of at least one of these is required for productive adhesion to P‐selectin (Pouyani and Seed, 1995; Wilkins et al., 1995). In CXCR‐3, CXCR‐4, and CCR‐2, tyrosine sulfation enhances ligand binding (Preobrazhensky et al., 2000; Fong et al., 2002; Veldkamp et al., 2006), while binding of HIV gp120 requires tyrosine sulfate on CCR‐5 for productive docking and viral entry (Farzan et al., 1999). A set of antibodies known as CD4i antibodies with extremely broad HIV recognition features a conspicuous and unusual sulfotyrosine in one of its hypervariable loops, which interact with a highly conserved arginine residue in HIV gp120 (Choe et al., 2003). Although, at least in these cases, sulfotyrosine‐based sulfotopes have critical biological roles, tyrosine sulfation does not appear to be actively regulated at the level of the tyrosine STs TPST‐1 and TPST‐2. These enzymes are broadly and constitutively expressed in almost all mammalian cells and appear to act indiscriminately on substrates with tyrosines located within specific consensus sequences (Moore, 2003).

5

Sulfation of O‐linked Glycans in Leukocyte Adhesion and Inflammation

5.1 Sulfated Glycans in Lymphocyte Recirculation Lymphocyte recirculation through peripheral lymphoid organs is a physiological process fundamental to host defense and immune surveillance. During this process, blood‐borne naı¨ve lymphocytes roll along and then firmly adhere to and transmigrate through the endothelium in specialized postcapillary vessels known as high endothelial venules (HEVs). In the lymph node parenchyma, the lymphocyte may encounter specific antigen, and hence proliferate and differentiate in germinal centers, most often however, the lymphocyte emigrates from the lymph node through the afferent lymph and recirculates to the blood. The first step within the multistep process of lymphocyte extravasation through HEVs involves tethering and rolling of the lymphocytes to and along the HEV endothelium and depends on the leukocyte‐homing receptor L‐selectin (Springer, 1994). L‐selectin binds to a set of highly glycosylated HEV ligands collectively known as peripheral lymph node addressin (PNAd) (Berg et al., 1991; Rosen, 2004). A unique characteristic of HEV is the capacity of its endothelial cells to incorporate large amounts of inorganic sulfate, both in human and in rodents (Andrews et al., 1982). Rosen and coworkers were the first to show that the mucin‐ type glycoproteins comprising PNad are highly sulfated (Imai et al., 1991) and that sulfation of PNAd was required to support binding of L‐selectin‐expressing lymphocytes to HEV (Imai et al., 1993). Further investigation defined the essential component of PNAd as the tetrasaccharide sialyl Lewis x (sLex) with a conspicuous 6‐O‐sulfation at its N‐acetylglucosamine (GlcNAc) moiety (Hemmerich and Rosen, 1994). This 6‐sulfo sLex tetrasaccharide caps complex O‐linked glycans that in their simplest form have the sulfotope connected through an extended core 1 linkage (Yeh et al., 2001) and/or a core 2 linkage (Hemmerich et al., 1995) to N‐acetylgalactosamine (GalNAc), which in turn is linked at its reducing end to a serine or threonine within the mucin core of the carrier protein (> Figure 9‐5).

5.2 Sulfated Glycans in Chronic Inflammation Leukocyte recruitment is also a hallmark of inflammation, and chronic inflammation appears to involve elaboration of lymphoid architecture in the inflamed tissue (Ruddle, 1999). This process includes extralymphoid differentiation of postcapillary vessels into HEV‐type vessels and/or de novo elaboration of the latter (Girard and Springer, 1995). These extralymphoid HEVs feature high‐affinity L‐selectin ligands similar to those expressed on lymph node HEVs, and facilitate extravasation of blood‐borne naı¨ve (bystander)

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. Figure 9‐5 Structure of the most simple O‐linked glycans on the L‐selectin PNAd. The GlcNAc6S on either branch is strictly required for high affinity L‐selectin binding and its biosynthesis is catalyzed by an ST expressed in the Golgi compartment of HEV endothelial cells. The substructures recognized by L‐selectin and the adhesion‐blocking sulfation‐dependent antibody MECA‐79 are highlighted

lymphocytes to the lesion (Rosen, 1999; van Zante and Rosen, 2003). As the chemokine repertoire in the inflammatory lesion differs from that in lymph node HEVs, these de novo HEV ligands can also recruit neutrophils and monocytes, both of which also express L‐selectin (Tedder et al., 1990). The investigation of extralymphoid L‐selectin ligands in settings of chronic inflammation has been greatly facilitated by the availability of a specific monoclonal antibody known as MECA‐79, which is adhesion‐blocking (Streeter et al., 1988) and sulfation dependent (Hemmerich et al., 1994), and was shown to specifically recognize the 6‐O‐sulfated N‐acetyllactosamine (Galb1!4GlcNAc6S) within the context of the core 1 extension (Yeh et al., 2001), a substructure contained in HEV ligands for L‐selectin (> Figure 9‐4). Using this reagent, a large number of studies have demonstrated the induction of sulfated extralymphoid L‐selectin ligands in many diseases associated with chronic inflammation in human and animal models, including rheumatoid arthritis (RA) in humans (Middleton et al., 2005; Pablos et al., 2005), collagen‐induced arthritis (CIA) in the B10RIII mouse (Yang et al., 2006), autoimmune diabetes in the NOD mouse (Hanninen et al., 1993), bronchial asthma in humans (Toppila et al., 2000) and in a sheep model (Rosen et al., 2005), inflammatory bowel disease (Middleton et al., 2005), and chronic heart and kidney allograft rejection (Toppila et al., 1999; Kirveskari et al., 2000).

5.3 Endothelial Sulfotransferases Since defining GlcNAc 6‐O‐sulfate (GlcNAc6S) as the critical determinant for L‐selectin binding, the identity of the ST(s) involved in L‐selectin ligand biosynthesis was pursued by a number of groups. Within the human ST complement only five of the eight enzymes in the galactose/GalNAc/GlcNAc 6‐O‐ST (GST) family are able to catalyze the 6‐O‐sulfation found in L‐selectin ligands, and of those only GST‐2 and GST‐3 are expressed in endothelial cells (Uchimura et al., 1998; Bistrup et al., 1999; Hiraoka et al., 1999), the latter being remarkably restricted to high endothelial cells (Bistrup et al., 1999; Hiraoka et al., 1999). Through analysis of GST‐deficient mice generated by gene targeting, GST‐3 was found to be responsible for the

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sulfation of a major portion of HEV‐ligands in peripheral lymph nodes (Hemmerich et al., 2001), specifically on the luminal aspect of HEV, while GST‐2 catalyzes sulfation of a minor portion of L‐selectin ligands (Uchimura et al., 2004), restricted mainly to the abluminal aspect of HEV. Deletion of both STs in double‐null mice resulted in complete absence of MECA‐79‐reactive high‐affinity HEV ligands (Kawashima et al., 2005; Uchimura et al., 2005); however, in vivo homing to lymph nodes still occurred at 30% of that seen in wild‐type mice, presumedly through compensation by unsulfated, low‐affinity L‐selectin ligands elaborated in the null animals. Deletion of GST‐3 also resulted in the absence of extralymphoid L‐selectin ligands from pancreatic islets in a transgenic mouse model of insulitis (Bistrup et al., 2004).

5.4 Induction of Endothelial Sulfotransferases in Chronic Inflammation As discussed above, sulfated extralymphoid L‐selectin ligands become induced on HEV‐like vessels at sites of chronic inflammation. As a number of sialomucins including CD34 and podocalyxin can function as L‐ selectin ligands (van Zante and Rosen, 2003), regulation of these ligands was expected to occur at the level of posttranslational modification (glycosylation and/or sulfation) rather than ligand transcription. In fact, CD34 is expressed on many types of endothelial cells and hematopoietic cells, however, only on lymphoid HEV (Satomaa et al., 2002) or extralymphoid HEV‐like vessels (Renkonen et al., 2002) is it decorated with the 6‐sulfo sLex sulfotope recognized by L‐selectin. Therefore regulation likely occurs at the level of glycan biosynthesis. Because of the requirement of GlcNAc 6‐O‐sulfation for L‐selectin binding, expression of the responsible STs GST‐3 and, to a lesser degree, GST‐2 is likely to regulate the elaboration of PNAd on HEVs and extralymphoid HEV‐like vessels at sites of chronic inflammation. Using enzyme‐specific antibodies, expression of GST‐3 was shown to correlate closely with PNAd expression in lymph node HEVs as well as extralymphoid vessels in chronically inflamed tissue but not healthy control tissue in human RA (Pablos et al., 2005) as well as in a number of animal models including murine CIA (Yang et al., 2006), sheep bronchial asthma (Rosen et al., 2005), and a transgenic mouse model of insulitis (Bistrup et al., 2004). In the CIA model, this induction of GST‐3 on the protein level is accompanied by a significant multifold increase in levels of transcripts encoding GST‐3 and to a lesser degree GST‐2 (Yang et al., 2006). While these data unambiguously show that de novo elaboration of PNAd occurs in many settings of chronic inflammation and is dependent on induction of the GlcNAc 6‐O‐STs GST‐3 and GST‐2, they do not prove a causative link between this phenomenon and the disease process. Rosen and coworkers (2005) recently provided initial evidence for the protective effect of the GlcNAc6S‐specific antibody MECA‐79 in a sheep model of asthma. In order to further prove (or disprove) this important hypothetical link between induction of GlcNAc 6‐O‐STs and chronic inflammation, GlcNAc 6‐O‐ST‐deficient mice are currently being bred into suitable backgrounds for investigation of phenotype in inflammatory models. If extralymphoid PNAd was shown critical to the disease process, it may emerge as an important target for broad spectrum antiinflammatory therapy, most likely through therapeutic IgG‐type antibodies against GlcNAc‐ 6‐sulfate in the context of PNAd‐derived sulfotopes. Moreover, the STs GST‐2 and GST‐3 will be targeted by small‐molecule inhibitors for indirect inhibition of PNAd function through inhibition of biosynthesis of its critical sulfotopes.

5.5 Other Immunologically Important Sulfotopes Sulfation of glycoproteins is rapidly emerging as an important immune‐modulatory mechanism. In > Sect. 3, I reviewed the role of tyrosine sulfation of the adhesion molecule PSGL‐1 as well as certain chemokine receptors involved in immune modulation and viral infection. Interestingly, in very specific situations, PSGL‐1 can carry glycan‐associated sulfotopes in addition to its tyrosine sulfation. Thus, a major subset of human dendritic cells, defined by the monoclonal antibody M‐DC8, carry a glycoform of PSGL‐1 in which the sLex tetrasaccharide is replaced by a 6‐sulfo‐lactosamine moiety (Schakel et al., 2002). Because of the lacking sLex, the DC‐associated PSGL‐1 glycoform does not bind to any of the selectins. However, the 6‐ sulfolactosamine elaborated on PSGL‐1 on M‐DC8þ dendritic cells is suspected to contribute to their

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pronounced proinflammatory capacity, probably through a not yet elucidated receptor‐mediated mechanism. Another glycoform of PSGL‐1 is expressed on TH2 cells, which also express high levels of CD45RO. In these cells at least a subset of PSGL‐1 molecules appears to be decorated with 6‐sulfo sLex rather than plain sLex (Ohmori et al., 2006) As 6‐sulfo sLex comprises the L‐selectin binding epitope on sulfoadhesin expressed on HEVs or HEV‐like vessels, these 6‐sulfo sLex positive TH2 cells bind as well to L‐selectin as they do to P‐selectin. The 6‐sulfo sLex determinant, in the context of PSGL‐1, was proposed to comprise at least part of the cutaneous lymphocyte antigen expressed on skin‐homing helper memory T cells (Fuhlbrigge et al., 1997; Ohmori et al., 2006).

6

Heparan Sulfate Proteoglycans

6.1 Composition and Biosynthesis of Heparan Sulfate Heparan sulfate (HS) proteoglycans (HSPGs) are glycoconjugates in which long linear chains of GlcNAc linked to glucuronic acid (GlcNAca1!4GlcAb1!4), also known as glucosaminoglycans (GAGs), are linked to a core protein such as syndecan, perlecan, or glypican (Varki and Chrispeels, 1999). These proteoglycans are anchored to the plasma membrane through their core protein or are secreted into the extracellular matrix. While their carbohydrate sequences are rather uniform, these GAGs carry very diverse and conspicuous sulfation modifications (> Figure 9‐1), which occur in sequence catalyzed by a large family of structurally related heparan sulfate N‐ and O‐STs. Furthermore, many of the GlcA residues within these chains become converted into iduronate (IdoA) residues through the action of a specific GlcA isomerase.

6.2 Biological Roles of HS‐Associated Sulfotopes The sulfation modifications in HSPGs occur in clusters and comprise diverse and often overlapping sulfotopes, which function as binding sites for a large variety of growth and differentiation factors, adhesion molecules, and chemoattractants. Heparin is a distinct, highly sulfated secreted HSPG produced by mast cells that along with other sulfotopes carries many copies of a multisulfated pentasaccharide defined as the antithrombin‐binding site. Binding of this pentasaccharide to antithrombin, and downstream inhibition of blood clotting by heparin depends on a specific GlcNAc 3‐O‐sulfation at the center of the hexasaccharide (Bjork and Lindahl, 1982). Arixtra (fondaparinux sodium) is a completely synthetic version of this sulfotope, now being marketed for antithrombotic therapy (Tan and Lip, 2005). Similar GlcNAc 3‐O‐ sulfate (GlcNAc3S)‐based sulfotopes were shown to be important viral entry receptors in herpes simplex (Shukla et al., 1999) and potentially hepatitis C (Cocquerel et al., 2006). Other sulfotopes that depend more on GlcNAc 6‐O‐sulfation mediate binding of growth factors such as vascular endothelial growth factor (VEGF) (Ono et al., 1999), platelet‐derived growth factor (PDGF) (Feyzi et al., 1997) and a number of fibroblast growth factors (FGFs) (Kreuger et al., 2001; Ashikari‐Hada et al., 2004), as well as adhesion molecules such as P‐ and L‐selectin (Wang et al., 2002), to HSPGs. The following subsection focuses on an important trisulfated disaccharide sulfotope on HSPGs that is important in development and cancer angiogenesis and is regulated by a novel class of extracellular endosulfatases.

6.3 Endosulfatases Involved in HS Remodeling The first of these enzymes, Q‐SULF, was discovered by Emerson and coworkers and was shown to link sonic hedgehog to Wnt signaling in myocyte development in the quail (Dhoot et al., 2001). Two mammalian homologues termed SULF‐1 and SULF‐2 were subsequently identified in humans and mice (Morimoto‐ Tomita et al., 2002). Both enzymes were expressed in CHO cells and were shown to hydrolyze a sulfate moiety linked to the 6‐position of GlcNAc in the trisulfated disaccharide fragment of HS, shown in > Figure 9‐6.

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. Figure 9‐6 Structure of the HS disaccharide structure desulfated at the 6‐position of its GlcNAc by the endosulfatases SULF1 and SULF2

Moreover, these enzymes are both able to release growth factors such as VEGF and FGF‐1 as well as chemokines SDF‐1 and SLC from immobilized heparin (Morimoto‐Tomita et al., 2002). Hanahan and coworkers had previously proposed a role of HSPGs in the angiogenic switch, a mechanism by which growing cancers induce proliferation of surrounding endothelium to gain access to blood supply. HS retains endothelial growth factors such as VEGF in an inactive, sequestered state, and degradation of HS by bacterial heparinases was shown to promote angiogenesis in a transgenic mouse model (Bergers et al., 2000). In mammals, there are no extracellular heparinases, rather, it is likely that the extracellular sulfatases SULF‐1 and SULF‐2 play key roles in the mobilization of endothelial growth factors. In fact, recombinant SULF‐2 induced vascular proliferation in the chick chorioallantoic membrane (CAM) assay, while an inactive mutant of SULF‐2 used as a control did not (Morimoto‐Tomita et al., 2002). Furthermore, SULF‐1 and SULF‐2 were shown to be upregulated in a number of human cancers including breast tumors (Morimoto‐Tomita et al., 2002) and pancreatic tumors (Steven D. Rosen and colleagues, unpublished). These findings prompted the working hypothesis illustrated in > Figure 9‐7. Endothelial cells and/or other parenchymal somatic cells express HSPGs that bind to endothelial growth factors and thus prevent them from binding to endothelial growth factor receptors. The growing cancer now secretes SULF‐1 and/or SULF‐2, which in turn hydrolyze critical 6‐O‐sulfates on GlcNAc within the HSPG‐associated growth factor‐binding sulfotopes, and therefore release the growth factor, which can now bind to its receptor on endothelial cells and thus promote tumor growth in an autocrine and/or paracrine fashion. While this hypothesis has not been proven unambiguously to date, it points to the angiogenic sulfatases SULF‐1 and SULF‐2 as attractive targets for novel cancer therapies.

7

Sulfation in Regulation of Hormone Half‐Life

The mammalian pituitary gland produces a plethora of hormones, which mediate endocrinal regulation of fundamental biological processes such as growth, development, metabolism, and reproduction. Two of the pituitary hormones, lutropin (LH) and thyrotropin (TSH), were found to bear asparagine‐linked glycans capped with GalNAc‐4‐O‐sulfate (GalNAc4S) moieties (Green et al., 1985; Green and Baenziger, 1988). These sulfated structures have been conserved through evolution and are found on glycoprotein hormones in all vertebrates from fish to humans (Manzella et al., 1995). The terminal GalNAc4S does not have any impact on hormone activity, rather it determines the circulatory half‐life of the hormone, because the sulfotope is recognized by the endocytic mannose/GalNAc4S receptor expressed on endothelial cells lining

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. Figure 9‐7 Hypothetical mechanism for the angiogenic switch in tumorigenesis—Angiogenic growth factors, specifically VEGF, are being maintained in an inactive sequestered state by sulfation‐dependent binding to HSPGs. SULF1 and/or SULF2 hydrolyze critical sulfate groups, and thus release VEGF, which can now promote endothelial proliferation by binding to a VEGF receptor on endothelial cells

the sinusoids of the liver. Native LH bearing the GlcNAc4S sulfotope is thus cleared from the circulation about fivefold more rapidly than recombinant LH, in which the N‐linked glycans are capped by sialylated lactosamine rather than GlcNAc4S (Baenziger et al., 1992). The shortened circulatory half‐life of native LH reduces its potency in vivo, but it is essential for obtaining a pulsatile rise and fall in LH level, which prevents desensitization of the LH receptor during the preovulatory surge of LH in the blood (Hooper et al., 1996). This GalNAc4S‐mediated regulation of LH circulatory half‐life was shown to be critical for implantation in vivo (Mi et al., 2002). The GalNAc4S sulfotope is generated through the sequential action of a ProXaaArg/Lys‐specific b1!4GalNAc transferase and GalNAc‐4‐O‐ST 1 (Dharmesh et al., 1993; Xia et al., 2000). GalNAc‐4‐O‐ST 1 is expressed in a number of different regions in the brain, predominantly in the pituitary, cerebellum, and pons (Baenziger, 2003). Nonhormone glycoproteins decorated with the GalNAc4S sulfotope were indeed detected in the cerebellum and are closely associated with Purkinje cells, the dominant type of neurons in this brain region (Nakayama et al., 1999). However, the function of the sulfotope in this setting is unclear.

8

Sulfation in the Central Nervous System

8.1 Sulfated Glycolipids in Myelin Structure and Function The central nervous system (CNS) is particularly rich in sulfated glycoproteins, proteoglycans, and specifically sulfated glycolipids. A prominent glycolipid in the CNS is sulfatide, a 3‐O‐sulfated galactosyl‐ ceramide (> Figure 9‐8). Sulfatide is a major component of the myelin sheet (4% of total lipid content) and is synthesized in oligodendrocytes (in the CNS) or Schwann cells (in the peripheral nervous system) (Ishizuka, 1997). Synthesis of sulfatide from its precursor is catalyzed by cerebroside ST (CST) (Honke et al., 1997), a member of the galactose 3‐O‐ST family of enzymes. Deletion of CST by gene targeting in mice results in complete absence of sulfatide in the brain (Honke et al., 2002). These animals are born healthy, but develop serious defects in neuronal conductance though their axons are well myelinated. However, these mice exhibit disorganized termination of the lateral loops at the nodes of Ranvier, and nodal clustering of Naþ and Kþ channels that generate the action potentials is also impaired (Ishibashi et al., 2002). Although the exact role of sulfatide in myelin organization and function remains unclear, it is

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. Figure 9‐8 Structure of sulfatide, a sulfated glycolipid and major component of CNS myelin

believed to play a critical role in regulating oligodendrocyte terminal differentiation (Honke et al., 2004), in addition to potential other roles as being a structural component of mature myelin. Sulfatide may also play an important role in multiple sclerosis (MS), a common autoimmune disorder characterized by progressive degradation of CNS myelin. In experimental autoimmune encephalitis (EAE), a mouse model of MS, leukocytes infiltrate the CNS, bind to myelin, and contribute to myelin degradation. Huang and coworkers (Huang et al., 1991, 1994) found that CNS myelin expresses ligands for L‐selectin. At about the same time, sulfatide was shown to support L‐selectin‐dependent leukocyte adhesion and rolling (Suzuki et al., 1993; Alon et al., 1995). Later, L‐selectin‐deficient mice were shown to be protected from paralysis in EAE even though leukocytes still infiltrated the CNS of these animals in the model (Grewal et al., 2001). Finally, in a mouse model of interstitial kidney inflammation, monocyte infiltration of the kidney was shown to be L‐ selectin dependent and considerably reduced in CST‐null mice that lack sulfatide (Ogawa et al., 2004). This set of data encourages speculation that myelin‐associated sulfatide supports leukocyte adhesion to myelin and its subsequent degradation by the leukocytes. Therefore, masking of the Gal‐3‐O‐sulfate‐dependent sulfotope in sulfatide by a specific therapeutic antibody or modulating its biosynthesis by CST inhibitors may afford therapeutic benefit for MS patients.

8.2 Chondroitin Sulfate in CNS Injury The CNS is also rich in proteoglycans, specifically chondroitin sulfate proteoglycans (CSPGs). Chondroitin sulfate (CS) is a linear polymer of GlcAb1!3GalNAcb1!4 with specific sulfation patterns, which allow definition of five subtypes: A, GlcA‐GalNAc4S; B, IdoA2S‐GalNAc4S; C, GlcA‐GalNAc6S; D, GlcA2S‐ GalNAc6S; and E, GlcA‐GalNAc4,6diS (Varki and Chrispeels, 1999). CSPGs serve as structural components of the CNS; in addition, CSPGs have also been implicated as negative regulators of neurite and axon growth (Hynds and Snow, 1999; Walz et al., 2002). More importantly, a number of CSPGs, such as aggrecan, neurocan, brevican, phosphacan, tenascin, and NG2, have been shown to contribute significantly to arrest of axon regeneration in the injured CNS (Sandvig et al., 2004). In a mouse model of spinal chord injury, treatment of the lesion with bacterial chondroitinase resulted in recovery of significant locomotor function in the injured animals (Bradbury et al., 2002). As there are no intrinsic extracellular chondroitinases in the mammalian CNS, the particular sulfation patterns found on CPSGs may be involved in the regulation of CPSG‐mediated inhibition of axon regeneration. Thus, in a rat model of brain injury, chondroitin 6‐O‐ST 1 was found upregulated in most glial cell types around cortical injuries, along with considerable increase of its product CS‐C (Properzi et al., 2005). Furthermore, CS‐E, a rather rare subtype of chondroitin sulfate, was also found differentially overexpressed in astroglial scars, and functions as a potent inhibitor of neurite extension (Gilbert et al., 2005). Formation of CS‐E requires the presence of GalNAc4S 6‐O‐ST (Ohtake et al., 2001), a yet poorly characterized enzyme originally discovered as a protein coexpressed with B‐rag

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and speculated to play an important role in B‐cell development (Verkoczy et al., 1998). Therefore, modulation of sulfation in CSPGs by inhibitors of the appropriate STs may be beneficial to promote axon regeneration in settings of brain or spinal cord injury.

8.3 Heparan Sulfate Proteoglycans in Axon Guidance and Targeting While CSPGs are typically thought of as inhibitory molecules that form barriers to axon growth, CNS‐ associated HSPGs are believed to be important in axon guidance and targeting (Van Vactor et al., 2006). HSPGs in neural development are typically expressed as regional gradients presenting distinct sulfotopes that direct axon growth, at least in part by modulating the response of the navigating growth cone to Slit proteins (Pratt et al., 2006). Structural features of these sulfotopes include IdoA 2‐O‐sulfation and GlcNAc‐ 6‐O‐sulfation (Irie et al., 2002). These HS sequences, rather than overall composition, appear to be essential for axon targeting and suggest that regionalized expression of specific HS‐STs is important for the creation of the guiding HSPG gradients.

8.4 Summary In summary, sulfated glycoconjugates play important roles in CNS structure, development, plasticity, and regeneration. Many potentially important sulfotopes on CNS‐associated glycoconjugates remain poorly defined structurally and functionally. Further elucidation of their biological roles and definition of the STs and sulfatases underlying their regulation may help toward a better understanding of CNS biology and potential novel therapies for yet intractable CNS disorders.

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Conclusions

Sulfation of glycoconjugates (glycoproteins, glycolipids, and proteoglycans) is a posttranslational modification fundamental to developmental biology, immunology, and neurobiology, as well as disease processes in cancer, inflammation, and CNS disorders. The enzymes at the heart of biological sulfation are the Golgi compartment‐associated STs which decorate their substrates with specific sulfated epitopes (sulfotopes), which eventually become exposed to the extracellular environment and comprise novel‐binding sites for biological effector molecules facilitating cell–cell and cell–matrix communications. Regulation of these sulfotopes occurs often at the level of the underlying sulfotransferase(s), and in some cases also by specific endosulfatases. Targeting of diseases processes involving specific sulfotopes can occur by specific antibodies to these sulfotopes or by small‐molecule inhibitors of Golgi compartment STs. In the latter case, because of certain structural and mechanistic similarities between sulfotransferses and kinases, lessons learned in kinase inhibitor design may be applicable to sulfotransferase inhibitor discovery.

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Protein sulfation Bergers G, et al. 2000. Matrix metalloproteinase‐9 triggers the angiogenic switch during carcinogenesis. Nat Cell Biol 2: 737-744. Bettelheim FR. 1954. Tyrosine‐O‐sulfate in a peptide from fibrinogen. J Am Chem Soc 76. Bick JA, Leustek T. 1998. Plant sulfur metabolism—the reduction of sulfate to sulfite. Curr Opin Plant Biol 1: 240-244. Bistrup A, et al. 1999. Sulfotransferases of two specificities function in the reconstitution of high endothelial cell ligands for L‐selectin. J Cell Biol 145: 899-910. Bistrup A, et al. 2004. Detection of a sulfotransferase (HEC‐ GlcNAc6ST) in high endothelial venules of lymph nodes and in high endothelial venule‐like vessels within ectopic lymphoid aggregates: Relationship to the MECA‐79 epitope. Am J Pathol 164: 1635-1644. Bjork I, Lindahl U. 1982. Mechanism of the anticoagulant action of heparin. Mol Cell Biochem 48: 161-182. Bradbury EJ, et al. 2002. Chondroitinase ABC promotes functional recovery after spinal cord injury. Nature 416: 636-640. Choe H, et al. 2003. Tyrosine sulfation of human antibodies contributes to recognition of the CCR5 binding region of HIV‐1 gp120. Cell 114: 161-170. Cocquerel L, et al. 2006. Hepatitis C virus entry: Potential receptors and their biological functions. J Gen Virol 87: 1075-1084. Dharmesh SM, et al. 1993. Co‐ordinate and restricted expression of the ProXaaArg/Lys‐specific GalNAc‐transferase and the GalNAcb1!4GlcNAcb1!2Mana 4‐sulfotransferase. J Biol Chem 268: 17096-17102. Dhoot GK, et al. 2001. Regulation of Wnt signaling and embryo patterning by an extracellular sulfatase. Science 293: 1663-1666. Falany CN. 1997. Enzymology of human cytosolic sulfotransferases. FASEB J 11: 206-216. Farzan M, et al. 1999. Tyrosine sulfation of the amino terminus of CCR5 facilitates HIV‐1 entry. Cell 96: 667-676. Feyzi E, et al. 1997. Characterization of heparin and heparan sulfate domains binding to the long splice variant of platelet‐ derived growth factor A chain. J Biol Chem 272: 5518-5524. Fong AM, et al. 2002. CX3CR1 tyrosine sulfation enhances fractalkine‐induced cell adhesion. J Biol Chem 277: 1941819423. Freeman C, Hopwood J. 1992. Lysosomal degradation of heparin and heparan sulphate. Adv Exp Med Biol 313: 121-134. Fuhlbrigge RC, et al. 1997. Cutaneous lymphocyte antigen is a specialized form of PSGL‐1 expressed on skin‐homing T cells. Nature 389: 978-981. Fukuda M, et al. 2001. Carbohydrate‐modifying sulfotransferases: Structure, function, and pathophysiology. J Biol Chem 276: 47747-47750. Gamage N, et al. 2006. Human sulfotransferases and their role in chemical metabolism. Toxicol Sci 90: 5-22.

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Gilbert RJ, et al. 2005. CS‐4,6 is differentially upregulated in glial scar and is a potent inhibitor of neurite extension. Mol Cell Neurosci 29: 545-558. Girard JP, Springer TA. 1995. High endothelial venules (HEVs): Specialized endothelium for lymphocyte migration. Immunol Today 16: 449-457. Girard JP, et al. 1999. Molecular cloning and functional analysis of SUT‐1, a sulfate transporter from human high endothelial venules. Proc Natl Acad Sci USA 96: 12772-12777. Green ED, Baenziger JU. 1988. Asparagine‐linked oligosaccharides on lutropin, follitropin, and thyrotropin. I. Structural elucidation of the sulfated and sialylated oligosaccharides on bovine, ovine, and human pituitary glycoprotein hormones. J Biol Chem 263: 25-35. Green ED, et al. 1985. Structural elucidation of the disulfated oligosaccharide from bovine lutropin. J Biol Chem 260: 15623-15630. Grewal IS, et al. 2001. CD62L is required on effector cells for local interactions in the CNS to cause myelin damage in experimental allergic encephalomyelitis. Immunity 14: 291-302. Hanninen A, et al. 1993. Vascular addressins are induced on islet vessels during insulitis in nonobese diabetic mice and are involved in lymphoid cell binding to islet endothelium. J Clin Invest 92: 2509-2515. Hashimoto Y, et al. 1992. Molecular cloning and expression of rat liver N‐heparan sulfate sulfotransferase. J Biol Chem 267: 15744-15750. Hemmerich S, Rosen SD. 1994. 60 ‐sulfated sialyl Lewis x is a major capping group of GlyCAM‐1. Biochemistry 33: 4830-4835. Hemmerich S, et al. 1994. Sulfation‐dependent recognition of high endothelial venules (HEV)‐ligands by L‐selectin and MECA 79, and adhesion‐blocking monoclonal antibody. J Exp Med 180: 2219-2226. Hemmerich S, et al. 1995. Structure of the O‐glycans in GlyCAM‐1, an endothelial‐derived ligand for L‐selectin. J Biol Chem 270: 12035-12047. Hemmerich S, et al. 2001. Sulfation of L‐selectin ligands by an HEV‐restricted sulfotransferase regulates lymphocyte homing to lymph nodes. Immunity 15: 237-247. Hemmerich S, et al. 2004. Strategies for drug discovery by targeting sulfation pathways. Drug Discov Today 9: 967-975. Hiraoka N, et al. 1999. A novel, high endothelial venule‐ specific sulfotransferase expresses 6‐sulfo sialyl Lewis x, an L‐selectin ligand displayed by CD34. Immunity 11: 79-89. Honke K, et al. 1997. Molecular cloning and expression of cDNA encoding human 30 ‐phosphoadenylylsulfate: Galactosylceramide 30 ‐sulfotransferase. J Biol Chem 272: 48644868. Honke K, et al. 2002. Paranodal junction formation and spermatogenesis require sulfoglycolipids. Proc Natl Acad Sci USA 99: 4227-4232.

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Honke K, et al. 2004. Biological roles of sulfoglycolipids and pathophysiology of their deficiency. Glycoconj J 21: 59-62. Hooper LV, et al. 1996. From legumes to leukocytes: Biological roles for sulfated carbohydrates. FASEB J 10: 1137-1146. Huang K, et al. 1991. A lymphocyte homing receptor (L‐ selectin) mediates the in vitro attachment of lymphocytes to myelinated tracts of the central nervous system. J Clin Invest 88: 1778-1783. Huang K, et al. 1994. Myelin localization of a central nervous system ligand for L‐selectin. J Neuroimmunol 53: 133-141. Hynds DL, Snow DM. 1999. Neurite outgrowth inhibition by chondroitin sulfate proteoglycan: Stalling/stopping exceeds turning in human neuroblastoma growth cones. Exp Neurol 160: 244-255. IHGSC—International Human Genome Sequencing Consortium 2001. Initial sequencing and analysis of the human genome. Nature 409: 860–921. IHGSC—International Human Genome Sequencing Consortium 2004. Finishing the euchromatic sequence of the human genome. Nature 431: 931–945. Imai Y, et al. 1991. Identification of a carbohydrate‐based endothelial ligand for a lymphocyte homing receptor. J Cell Biol 113: 1213-1221. Imai Y, et al. 1993. Sulfation requirement for GlyCAM‐1, an endothelial ligand for L‐selectin. Nature 361: 555-557. Irie A, et al. 2002. Specific heparan sulfate structures involved in retinal axon targeting. Development 129: 61-70. Ishibashi T, et al. 2002. A myelin galactolipid, sulfatide, is essential for maintenance of ion channels on myelinated axon but not essential for initial cluster formation. J Neurosci 22: 6507-6514. Ishizuka I. 1997. Chemistry and functional distribution of sulfoglycolipids. Prog Lipid Res 36: 245-319. Kakuta Y, et al. 1998a. Conserved structural motifs in the sulfotransferase family. Trends Biochem Sci 23: 129-130. Kakuta Y, et al. 1998b. The sulfuryl transfer mechanism. Crystal structure of a vanadate complex of estrogen sulfotransferase and mutational analysis. J Biol Chem 273: 27325-27330. Kakuta Y, et al. 1999. Crystal structure of the sulfotransferase domain of human heparan sulfate N‐deacetylase/N‐sulfotransferase 1. J Biol Chem 274: 10673-10676. Kamiyama S, et al. 2003. Molecular cloning and identification of 30 ‐phosphoadenosine 50 ‐phosphosulfate transporter. J Biol Chem 278: 25958-25963. Kauffman FC. 2004. Sulfonation in pharmacology and toxicology. Drug Metab Rev 36: 823-843. Kawashima H, et al. 2005. N‐acetylglucosamine‐6‐O‐sulfotransferases 1 and 2 cooperatively control lymphocyte homing through L‐selectin ligand biosynthesis in high endothelial venules. Nat Immunol 6: 1096-1104.

Kehoe JW, Bertozzi CR. 2000. Tyrosine sulfation: A modulator of extracellular protein–protein interactions. Chem Biol 7: R57-R61. Kirveskari J, et al. 2000. De novo induction of endothelial L‐ selectin ligands during kidney allograft rejection. J Am Soc Nephrol 11: 2358-2365. Kopriva S. 2006. Regulation of sulfate assimilation in Arabidopsis and beyond. Ann Bot (Lond) 97: 479-495. Kreuger J, et al. 2001. Sequence analysis of heparan sulfate epitopes with graded affinities for fibroblast growth factors 1 and 2. J Biol Chem 276: 30744-30752. Lukatela G, et al. 1998. Crystal structure of human arylsulfatase A: The aldehyde function and the metal ion at the active site suggest a novel mechanism for sulfate ester hydrolysis. Biochemistry 37: 3654-3664. Manzella SM, et al. 1995. Evolutionary conservation of the sulfated oligosaccharides on vertebrate glycoprotein hormones that control circulatory half‐life. J Biol Chem 270: 21665-21671. Markovich D. 2001. Physiological roles and regulation of mammalian sulfate transporters. Physiol Rev 81: 14991533. Meh DA, et al. 2001. The amino acid sequence in fibrin responsible for high affinity thrombin binding. Thromb Haemost 85: 470-474. Mi Y, et al. 2002. Regulation of lutropin circulatory half‐life by the mannose/N‐acetylgalactosamine‐4‐SO4 receptor is critical for implantation in vivo. J Clin Invest 109: 269-276. Middleton J, et al. 2005. A comparative study of endothelial cell markers expressed in chronically inflamed human tissues: MECA‐79, Duffy antigen receptor for chemokines, von Willebrand factor, CD31, CD34, CD105, and CD146. J Pathol 206: 260-268. Moore KL. 2003. The biology and enzymology of protein tyrosine O‐sulfation. J Biol Chem 278: 24243-24246. Morimoto‐Tomita M, et al. 2002. Cloning and characterization of two extracellular heparin‐degrading endosulfatases in mice and humans. J Biol Chem 277: 49175-49185. Mougous JD, et al. 2006. Molecular basis for G protein control of the prokaryotic ATP sulfurylase. Mol Cell 21: 109-122. Nakayama J, et al. 1999. Expression cloning of a human a1!4‐ N‐acetylglucosaminyltransferase that forms GlcNAca1!4Galb!R, a glycan specifically expressed in the gastric gland mucous cell‐type mucin. Proc Natl Acad Sci USA 96: 89918996. Ogawa D, et al. 2004. Cerebroside sulfotransferase deficiency ameliorates L‐selectin‐dependent monocyte infiltration in the kidney after ureteral obstruction. J Biol Chem 279: 2085-2090. Ohmori K, et al. 2006. Identification of cutaneous lymphocyte‐associated antigen as sialyl 6‐sulfo Lewis x, a selectin ligand expressed on a subset of skin‐homing helper memory T cells. Blood 107: 3197-3204.

Protein sulfation Ohtake S, et al. 2001. Human N‐acetylgalactosamine 4‐sulfate 6‐O‐sulfotransferase cDNA is related to human B cell recombination activating gene‐associated gene. J Biol Chem 276: 43894-43900. Ono K, et al. 1999. Structural features in heparin that interact with VEGF165 and modulate its biological activity. Glycobiology 9: 705-711. Pablos JL, et al. 2005. A HEV‐restricted sulfotransferase is expressed in rheumatoid arthritis synovium and is induced by lymphotoxin‐a/b and TNF‐a in cultured endothelial cells. BMC Immunol 6: 6. Page R. 2005. TreeView X: An open source program to display phylogenetic trees on Linux, Unix, Mac OS X, and Windows platforms. http://darwin.zoology.gla.ac.uk/rpage/treeviewx/ Pouyani T, Seed B. 1995. PSGL‐1 recognition of P‐selectin is controlled by a tyrosine sulfation consensus at the PSGL‐1 amino terminus. Cell 83: 333-343. Pratt T, et al. 2006. Heparan sulfation patterns generated by specific heparan sulfotransferase enzymes direct distinct aspects of retinal axon guidance at the optic chiasm. J Neurosci 26: 6911-6923. Preobrazhensky AA, et al. 2000. Monocyte chemotactic protein‐1 receptor CCR2B is a glycoprotein that has tyrosine sulfation in a conserved extracellular N‐terminal region. J Immunol 165: 5295-5303. Properzi F, et al. 2005. Chondroitin‐6‐sulphate synthesis is upregulated in injured CNS, induced by injury‐related cytokines, and enhanced in axon‐growth inhibitory glia. Eur J Neurosci 21: 378-390. Renkonen J, et al. 2002. Glycosylation might provide endothelial zip codes for organ‐specific leukocyte traffic into inflammatory sites. Am J Pathol 161: 543-550. Roeser D, et al. 2006. A general binding mechanism for all human sulfatases by the formylglycine‐generating enzyme. Proc Natl Acad Sci USA 103: 81-86. Rosen SD. 1999. Endothelial ligands for L‐selectin: From lymphocyte recirculation to allograft rejection. Am J Pathol 155: 1013-1020. Rosen SD. 2004. Ligands for L‐selectin: Homing, inflammation, and beyond. Annu Rev Immunol 22: 129-156. Rosen SD, et al. 2005. Therapeutic targeting of endothelial ligands for L‐selectin (PNAd) in a sheep model of asthma. Am J Pathol 166: 935-944. Ruddle NH. 1999. Lymphoid neoorganogenesis: Lymphotoxin’s role in inflammation and development. Immunol Res 19: 119-125. Sandvig A, et al. 2004. Myelin‐, reactive glia‐, and scar‐derived CNS axon growth inhibitors: Expression, receptor signaling, and correlation with axon regeneration. Glia 46: 225-251. Satomaa T, et al. 2002. O‐glycans on human high endothelial CD34 putatively participating in L‐selectin recognition. Blood 99: 2609-2611.

9

Schakel K, et al. 2002. 6‐Sulfo LacNAc, a novel carbohydrate modification of PSGL‐1, defines an inflammatory type of human dendritic cells. Immunity 17: 289-301. Segal IH, et al. 1987. Sulfate‐activating enzymes. Methods Enzymol 143: 334-350. Shukla D, et al. 1999. A novel role for 3‐O‐sulfated heparan sulfate in herpes simplex virus 1 entry. Cell 99: 13-22. Springer TA. 1994. Traffic signals for lymphocyte recirculation and leukocyte emigration: The multistep paradigm. Cell 76: 301-314. Streeter PR, et al. 1988. Immunohistologic and functional characterization of a vascular addressin involved in lymphocyte homing into peripheral lymph nodes. J Cell Biol 107: 1853-1862. Suzuki Y, et al. 1993. Sulfated glycolipids are ligands for a lymphocyte homing receptor, L‐selectin (LECAM‐1), binding epitope in sulfated sugar chain. Biochem Biophys Res Commun 190: 426-434. Tan KT, Lip GY. 2005. Fondaparinux. Curr Pharm Des 11: 415-419. Tedder TF, et al. 1990. Expression of the human leukocyte adhesion molecule, LAM1. Identity with the TQ1 and Leu‐ 8 differentiation antigens. J Immunol 144: 532-540. Thompson JD, et al. 1997. The CLUSTALX windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25: 4876-4882. Toppila S, et al. 1999. Endothelial L‐selectin ligands are likely to recruit lymphocytes into rejecting human heart transplants. Am J Pathol 155: 1303-1310. Toppila S, et al. 2000. Endothelial sulfated sialyl Lewis x glycans, putative L‐selectin ligands, are preferentially expressed in bronchial asthma but not in other chronic inflammatory lung diseases. Am J Respir Cell Mol Biol 23: 492-498. Uchimura K, et al. 1998. Molecular cloning and characterization of an N‐acetylglucosamine‐6‐O‐sulfotransferase. J Biol Chem 273: 22577-22583. Uchimura K, et al. 2004. N‐acetylglucosamine 6‐O‐sulfotransferase‐1 regulates expression of L‐selectin ligands and lymphocyte homing. J Biol Chem 279: 35001-35008. Uchimura K, et al. 2005. A major class of L‐selectin ligands is eliminated in mice deficient in two sulfotransferases expressed in high endothelial venules. Nat Immunol 6: 1105-1113. Van Vactor D, et al. 2006. Heparan sulfate proteoglycans and the emergence of neuronal connectivity. Curr Opin Neurobiol 16: 40-51. van Zante A, Rosen SD. 2003. Sulfated endothelial ligands for L‐selectin in lymphocyte homing and inflammation. Biochem Soc Trans 31: 313-317. Varki A, Chrispeels MJ. 1999. Essentials of Glycobiology. Cold Spring Harbor Laboratory Press.Cold Spring Harbor, N.Y.,

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Wang L, et al. 2002. Heparin’s antiinflammatory effects require glucosamine 6‐O‐sulfation and are mediated by blockade of L‐ and P‐selectins. J Clin Invest 110: 127-136. Wilkins PP, et al. 1995. Tyrosine sulfation of P‐selectin glycoprotein ligand‐1 is required for high affinity binding to P‐ selectin. J Biol Chem 270: 22677-22680. Xia G, et al. 2000. Molecular cloning and expression of the pituitary glycoprotein hormone N‐acetylgalactosamine‐4‐ O‐sulfotransferase. J Biol Chem 275: 38402-38409. Yang J, et al. 2006. Induction of PNAd and N‐acetylglucosamine 6‐O‐sulfotransferases 1 and 2 in mouse collagen‐induced arthritis. BMC Immunol 7: 12. Yeh JC, et al. 2001. Novel sulfated lymphocyte homing receptors and their control by a core1 extension b1!3‐N‐acetylglucosaminyltransferase. Cell 105: 957-969.

10

Protein Folding

A. Szila´gyi . J. Kardos . S. Osva´th . L. Barna . P. Za´vodszky

1 1.1

Protein Structure and Its Physical Basis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305 Physical Forces and Principles Underlying Protein Folding and Structure . . . . . . . . . . . . . . . . . . . . . 305

2

The Protein‐Folding Problem . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 306

3

Folding Mechanisms and Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 308

4 4.1 4.2

Molten Globules and Other Compact Denatured States . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310 The Structure of the Molten Globule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310 The Role of the Molten Globule in Folding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311

5 5.1 5.2 5.3

Simulations of Protein Folding and Unfolding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 312 All‐Atom Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 312 Simplified Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313 Multiscale Modeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313

6

Free‐Energy Landscapes of Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313

7 7.1 7.2

Traps on the Folding Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 315 Backbone Isomerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 315 Formation of Disulfide Bridges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 317

8

Transition States on the Folding Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 318

9 9.1 9.2

Folding of Multidomain and Multi‐Subunit Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320 Multidomain Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320 Multi‐Subunit Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321

10 10.1 10.2 10.3 10.4 10.5 10.6

Protein Folding in the Cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 The Hsp70 Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 323 The Folding Cage of Chaperonins (Hsp60 Family) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 324 The Hsp90 Chaperone System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325 Chaperone‐Assisted Assembly of Cellular Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325 Cooperation Between the Different Chaperone Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325 Chaperones and Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325

11 11.1 11.2 11.3 11.4 11.5 11.6

Protein Misfolding and Aggregation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 Degenerative Diseases Associated with Amyloid Deposition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 The Structure and Morphology of Amyloid Fibrils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 Mechanism of Amyloid Fibril Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 328 Amyloid Formation of Globular Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 328 Physicochemical and Sequence Determinants of Amyloid Formation . . . . . . . . . . . . . . . . . . . . . . . . . . 329 Factors Inducing or Inhibiting Amyloid Formation Under Physiological Conditions . . . . . . . . . 330

#


304

10

Protein folding

12 12.1 12.2 12.3 12.4 12.5 12.6

New Biophysical Techniques for the Study of Protein Folding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 330 Rapid Mixing Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 330 Real‐Time NMR Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 330 Chemically Induced Nuclear Polarization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 High‐Pressure NMR Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 Protein Folding and Dynamics Studied by Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332 Mechanical Unfolding of Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332

13

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332

Protein folding

10

Abstract: Since Anfinsen’s famous experiments in the 1960s, it has been known that the complex three‐ dimensional structure of protein molecules is encoded in their amino acid sequences, and the chains autonomously fold under proper conditions. Cracking this code, which is sometimes called ‘‘the second part of the genetic code,’’ has been one of the greatest challenges of molecular biology. Although a full understanding of how proteins fold remains elusive, theoretical and experimental studies of protein folding have come a long way since Anfinsen’s findings. In the living cell, folding occurs in a complex and crowded environment, often involving helper proteins, and in some cases it can go awry: the protein can misfold, aggregate, or form amyloid fibers. It is increasingly recognized that misfolded proteins and amyloid formation are the root cause of a number of serious illnesses including several neurodegenerative diseases. Therefore, the study of protein folding remains a key area of biomedical research. List of Abbreviations: AFM, atomic force microscopy; CCP, chaperonin‐containing TCP‐1; CIDNP, chemically induced nuclear polarization; DMD, discrete molecular dynamics; EM, electron microscopy; ESIMS, electrospray ionization mass spectrometry; FIS, factor for inversion stimulation; FMN, flavin mononucleotide; NOE, nuclear Overhauser effect; TF, trigger factor; TCP‐1, tailless complex polypeptide‐1

1

Protein Structure and Its Physical Basis

The function of a protein can only be interpreted from its structure. The nervous system is a network of cells, and the peculiar functional properties of these cells can be derived from the properties and interactions of their proteins. Proteins are involved in all stages of neural activity. Those embedded wholly or partly in membranes regulate the transport of ions and molecules as a means of signal exchange with other cells and the external medium. Some of them have enzymatic functions to catalyze the chemical processes essential for function. The diverse and highly specific function of proteins is a consequence of their sophisticated, individual surface pattern regarding shape, charge, and hydrophobicity. The surface pattern is a consequence of the unique three‐dimensional structure of the polypeptide chain. Proteins are linear polymers with nonrepetitive, specific covalent structure. The covalent structure is determined by the order of amino acids in which they are linked together. Since Anfinsen’s famous experiments (1973) in the 1960s, it has been believed and today generally accepted that folding and the resulting native structure of proteins are autonomously governed and determined by the amino acid sequence of a particular protein and its natural solvent environment.

1.1 Physical Forces and Principles Underlying Protein Folding and Structure A linear polypeptide chain is autonomously organized into a space‐filling, compact, and well‐defined three‐ dimensional structure. In a globular protein, the internal core is mostly formed by hydrophobic amino acid residues, held together by van der Waals forces, and the surface of the globule is formed by mostly charged and polar side chains. Proteins exist in this state of condensed matter while the specific conformation is largely determined by the flexibility of the polypeptide backbone and by the specific, consistent intermolecular interactions of the side chains. The monomeric unit in a polypeptide chain is the peptide group. The sequence of amino acids is the primary structure of the protein. The C, O, N, and H atoms lie in the same plane; successive planes define angles f and c. The conformation of a chain of n amino acids can be defined by 2n parameters. The restricted flexibility of the polypeptide chain is a major factor among those determining protein structure and folding. The native conformation must be energetically stable. From a thermodynamic point of view, the free energy of a protein molecule is influenced by the following major contributions: (1) the hydrophobic effect, (2) the energy of hydrogen bonds, (3) the energy of electrostatic interactions, and (4) the conformational entropy due to the restricted motion of the main chain and the side chains.

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The hydrophobic effect used to be explained as a primarily entropic effect arising from the rearrangement of hydrogen bonds between solvent molecules around an apolar solute. This hydration process is energetically unfavorable, and therefore drives apolar solutes together, thereby decreasing their solvent‐ exposed surface area. Today, the hydrophobic effect is usually viewed as a combined effect of hydration (an entropic effect) and van der Waals interactions between solute molecules (an enthalpic effect) (Makhatadze and Privalov, 1995). It is therefore entropic at low temperatures and enthalpic at high temperatures, which results in a complex temperature dependence of its strength (Schellman, 1997). Nevertheless, the hydrophobic force has long been considered as the major driving force of protein folding (Dill, 1990) as it leads to a rapid collapse of the polypeptide chain, thereby largely reducing the configurational space to explore. Without doubt, the hydrophobic interaction is also a major stabilizing force contributing to the thermodynamic stability of the folded state. The role of hydrogen bonds in folding and stability used to be underestimated based on the argument that intramolecular hydrogen bonds can be replaced by hydrogen bonds between the protein and the solvent. After a number of mutational studies, however, hydrogen bonds have now been recognized as having a contribution to protein stability as important as the hydrophobic effect (Pace et al., 1996). This contribution was estimated to be 1.5 1.0 kcal/mol per buried intramolecular hydrogen bond (Pace et al., 1996). Electrostatic interactions such as ion pairs and salt bridges in proteins have been an area of active research (Kumar and Nussinov, 2002). While hydrogen bonds and hydrophobic forces are essentially nonspecific, electrostatic interactions are largely specific, and therefore play an important role in specifying the fold of a protein as well as in protein flexibility and function. Computational and experimental evidence shows that salt bridges can be stabilizing or destabilizing. On the other hand, genome‐wide and structural comparisons of thermophilic and mesophilic proteins indicate that salt bridges may significantly contribute to the enhanced thermal stability of proteins from thermophilic organisms (Szilagyi and Zavodszky, 2000; Li et al., 2005; Razvi and Scholtz, 2006). The major destabilizing contribution to the stability of the folded state is the conformational entropy of the polypeptide chain. Folding a long chain into a specific, compact structure obviously results in a significant entropy decrease. This is counterbalanced by the various intrachain interactions described above. The resulting overall stability of the protein (the free‐energy difference between the folded and the unfolded state) is marginal, being on the order of 5–10 kcal/mol. This number is a small difference between huge stabilizing and destabilizing contributions. We qualitatively know that the hydrophobic effect and hydrogen bonds are the major stabilizing contributions and the conformational entropy is the major destabilizing one. However, due to the compensatory effects in the total energy balance, a quantitative prediction with respect to the significance of any specific type of interaction cannot be made with confidence (Jaenicke, 2000).

2

The Protein‐Folding Problem

The ‘‘central dogma’’ of molecular biology states that the flow of sequential information from nucleic acid to protein is unidirectional: nucleic acid sequences encode the sequence of proteins but once translation occurs, information cannot flow back from protein to nucleic acid. A possible extension of the central dogma would be to add that the sequence of the protein ‘‘encodes’’ its three‐dimensional structure. Indeed, this coding is sometimes called the ‘‘second half of the genetic code.’’ Cracking this code would be equivalent to solving the ‘‘protein folding problem.’’ To see what this problem is all about, let us look at Anfinsen’s classic experiment (1973) (> Figure 10-1). Ribonuclease, an enzyme with 124‐amino acid residues, contains four disulfide bridges. Treatment of ribonuclease with 8 M urea in the presence of the reducing agent b‐mercaptoethanol causes a complete unfolding of the ribonuclease molecule, yielding an essentially random form. In this process, the four disulfide bridges get cleaved, resulting in eight free SH groups. The enzymatic activity of the molecule is completely lost. Allowing the cysteines to reoxidize under denaturing conditions results in a mixture of ‘‘scrambled’’ species where the eight SH groups randomly pair to form four disulfide bridges (it is easy to

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. Figure 10-1 Scheme of Anfinsen’s experiment. See text for details

calculate that there are 105 possible pairings). However, when urea is slowly removed by dialysis and a small amount of b‐mercaptoethanol is added, disulfide interchange takes place and the mixture of ‘‘scrambled’’ ribonuclease molecules is eventually converted to a homogeneous product, which is fully enzymatically active and indistinguishable from native ribonuclease. This crucial experiment supports the idea known as the ‘‘thermodynamic hypothesis,’’ which states that the three‐dimensional structure of a native protein in its normal physiological state is the one in which the Gibbs free energy of the whole system is lowest. Therefore, in a given environment, the native conformation of the protein is fully determined by its amino acid sequence. How the information encoded in the sequence gets translated into the three‐dimensional structure? This is the protein‐folding problem. Assuming that the polypeptide chain randomly samples all possible configurations, we can estimate the time required for a protein to fold. If each bond connecting two neighboring amino acids can have, say, three possible states then a protein of 101 amino acids could exist in 3100 ¼ 51047 different configurations. Only one of these configurations corresponds to the native state. Even if the protein is able to sample new configurations at a rate of 1013 per second, it will take 1027 years to try them all. Nevertheless, proteins do fold, and in a timescale of seconds or less. This contradiction was first pointed out by Cyrus Levinthal in 1969 (Levinthal, 1969) and has become known as ‘‘Levinthal’s paradox.’’ To resolve the paradox, Levinthal argued that the protein cannot fold by random search and there must be specific ‘‘folding pathways.’’ The concept of folding pathways motivated a large number of experimental studies aimed at finding the specific ‘‘folding intermediates’’ and also gave rise to a number of models describing the folding process. For example, the nucleation/growth model (Wetlaufer, 1973) tried to resolve Levinthal’s paradox by assuming that the rate‐limiting step of the folding process is a nucleation event, presumably the formation of smaller structural units, and once nucleation occurs the nuclei grow fast and the folding process rapidly completes. This model is not consistent with the large number of observations where folding intermediates were observed. According to the ‘‘diffusion–collision–adhesion model’’ (Karplus and Weaver, 1976), fluctuating microdomains (portions of secondary structure or hydrophobic clusters) move diffusively and repeatedly collide with each other. Collisions can lead to a coalescence of the microdomains into larger units (adhesion). The rate‐limiting stage is assumed to be the diffusion process. This model is well supported by many experiments (Karplus and Weaver, 1994). The ‘‘framework model’’ (Baldwin, 1989) states that the

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folding process is hierarchical, starting with the formation of the secondary structure elements, and the docking of the preformed substructures is the rate‐limiting step. The ‘‘hydrophobic collapse model’’ (Dill, 1985) is based on the view that the hydrophobic effect is the main driving force of folding, and the process starts with a rapid collapse of the chain, followed by the formation of the secondary structure. In fact, whether hydrophobic collapse or secondary structure formation occurs first has remained a largely undecided issue even to this day. Finally, the ‘‘jigsaw puzzle model’’ (Harrison and Durbin, 1985) denied the necessity of a unique, directed folding pathway and stated that each protein molecule can follow a different route to the native structure, just like there are multiple ways to solve a jigsaw puzzle. This idea is actually consistent with a ‘‘new view’’ of protein folding, which gained popularity in the 1990s: the energy landscape view. The energy landscape view likens the energy landscape of a protein to a funnel, with the native structure at its global minimum, and each molecule may follow a different microscopic route from the top to the bottom (energy landscapes are discussed in more detail later). The many models of protein folding are not mutually exclusive; they try to grasp different aspects of folding, and experimental results give some support to each model we mentioned. A newer model, named ‘‘nucleation–condensation model,’’ is an attempt to unite the features of both the framework and the hydrophobic collapse mechanisms (Fersht, 1995; Fersht, 1997). In this model, long‐range and other native hydrophobic interactions form in the transition state to stabilize the otherwise weak secondary structure. The framework and the hydrophobic collapse models are viewed as two extremes of the nucleation–condensation mechanism; most proteins fold by a mechanism that is somewhere between the two extremes, i.e., secondary structure and hydrophobic interactions form nearly simultaneously and synergistically (Daggett and Fersht, 2003).

3

Folding Mechanisms and Kinetics

A unified view of protein folding should be general enough to interpret the diverse experimental findings of the field. Thermodynamics offers such a universal approach. Thermodynamic systems in equilibrium occupy the states with lowest Gibbs free energy at constant pressure and temperature. The Gibbs free energy (G) consists of an enthalpy and an entropic term G ðqÞ ¼ H ðqÞ T ðqÞ; where H is the enthalpy, T the absolute temperature, and S the entropy of the protein, and q represents the reaction coordinate used to describe the progress of the protein advancing from the unfolded toward the native state. Under physiological conditions, proteins maintain their native structure because the favorable enthalpic term arising from the solvent and protein interactions exceeds in magnitude the unfavorable entropic term, and therefore the native state has a smaller Gibbs free energy than the denatured state. The stability of the protein depends on the solvent–solvent, protein–solvent, and protein–protein interactions. These interactions depend on the intensive parameters that describe the thermodynamic state of the system. The enthalpic and the entropic terms are large, but of opposite sign, and almost cancel each other. The Gibbs free‐energy difference between the biologically active and denatured states of the proteins is rather small (Scharnagl et al., 2005). Proteins are stable only within a narrow range of conditions and can be denatured by changing virtually any of the intensive parameters (Shortle, 1996). Experiments prove that proteins can be unfolded by heat (Tsai et al., 2002; Prabhu and Sharp, 2005), cold (Franks, 1995; Kunugi and Tanaka, 2002), high pressure (Smeller, 2002; Meersman et al., 2006), extreme pH (Puett, 1973; Fitch et al., 2006), and addition of salts (Pfeil, 1981). Studies of protein stability and folding systematically change one or more of the intensive parameters and follow the kinetics of the change and/or the shift of equilibrium. There is a broad selection of methods that can be used to follow the structural changes of the proteins, including fluorescence (Isenman et al., 1979; Vanhove et al., 1998), phosphorescence (Mersol et al., 1993; Mazhul’ et al., 2003), circular dichroism (Kelly and Price, 2000), infrared spectroscopy (Fabian and Naumann, 2004; Ma and Gruebele, 2005),

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nuclear magnetic resonance (Englander and Mayne, 1992; Kamatari et al., 2004), and mass spectroscopy (Miranker et al., 1996; Konermann and Simmons, 2003). Both theoretical and experimental results indicate that a single reaction coordinate in general is not enough to describe protein folding, and multiple reaction coordinates must be used. (Becker and Karplus, 1997; Ma and Gruebele, 2005). Finding the adequate reaction coordinates for protein folding is not straightforward. Several kinetic and thermodynamic coordinates have been used to describe the ‘‘nativeness’’ of a given protein state. Thermodynamic reaction coordinates use a thermodynamic parameter, e.g., Gibbs free energy and/or entropy, to define the distance between the native state and the actual state of a protein. The kinetic reaction coordinate measures the time needed for the protein to reach the native state from a given starting state. An important thermodynamic reaction coordinate often used to describe the folding process is the number of native contacts present in the conformation, which proved useful in interpreting simple folding processes. Thermodynamic reaction coordinates, however, are inadequate to describe folding dominated by kinetic traps because they completely ignore the Gibbs free‐energy barriers separating the different states (Sali et al., 1994; Wolynes et al., 1995; Chan and Dill, 1998). The Gibbs free‐energy barrier to folding is determined by the unfavorable loss in configurational entropy upon folding and the gain in stabilizing native interactions. Starting from the unfolded protein, the polypeptide chain has to fold partially in order to bring together the residues that need to form the contacts stabilizing the native structure. The constrained polypeptide chain has smaller entropy, which means higher Gibbs free energy. As native contacts form, the enthalpy term decreases, the protein is stabilized. The rate‐limiting step in the folding process is the formation of the transition state, i.e., the conformation that has the highest Gibbs free energy on the folding pathway (Chan and Dill, 1998; Lindorff‐ Larsen et al., 2005). The simplest model for unfolding and refolding involves a single cooperative folding step, in which the unfolded (U) and folded (F) states of the protein interconvert: U ↔ F. This simple mechanism well describes the folding of several small proteins (Gillespie and Plaxco, 2004). The formation of a contact between two residues in the transition state involves an entropic cost which depends on the sequence separation of the two residues: the longer the chain between them the greater the entropic cost, and this entropic cost contributes to the height of the Gibbs free‐energy barrier between the unfolded and the folded state. If nonnative interactions play a marginal role in the transition state, it is possible to estimate the folding kinetics from the average sequence separation of the contacts in the native structure (Plaxco et al., 1998; Grantcharova et al., 2001; Zarrine‐Afsar et al., 2005). Intermediate structures were observed to accumulate during the folding of many proteins (Englander, 2000). Such intermediate states are trapped structures that have low Gibbs free energy. Mass action models that involve one or more intermediate states were constructed to explain more complex folding kinetics. Mass action models distinguish between ‘‘on‐pathway’’ and ‘‘off‐pathway’’ intermediates depending on whether the intermediate is on the folding pathway between the unfolded and native states (Baldwin, 1996). Off‐pathway intermediates often correspond to misfolded structures that must completely or partially unfold to allow formation of the native state (Evans et al., 2005). The most general theory of protein folding is a statistical mechanical model that uses the concept of the energy landscape, which is discussed in more detail later. Here we only want to clarify that in the energy landscape view, there is no clear distinction between on‐ and off‐ pathway intermediates. Folding mechanisms involving these two types of intermediates only differ in the distribution of traps on a Gibbs free‐energy landscape (Onuchic and Wolynes, 2004; Jahn and Radford, 2005). Energy landscape theory of protein folding predicts that the enthalpic and the entropic term of the transition‐state Gibbs free energy can cancel each other, leading to folding that lacks an activation barrier (Bryngelson et al., 1995). Although such downhill folding has indeed been found experimentally, it seems to be atypical, probably because such proteins are evolutionary unfavorable (Yang and Gruebele, 2004a). The probable reason for this is that proteins that fold downhill lack the barrier that prevents partial unfolding, and thus become more prone to aggregation (Yang and Gruebele, 2003; Gruebele, 2005). The folding process is usually not restricted to a narrow path in conformational space. Each molecule may follow a different path, and the molecule population can be very heterogeneous. Mass action models are inadequate to describe these folding processes, and the more complex energy landscape view must be used.

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Heterogeneous folding ensembles give rise to stretched folding kinetics and/or probe‐dependent observation of the kinetics (Sabelko et al., 1999; Osvath et al., 2003; Ma and Gruebele, 2005; Osvath et al., 2006).

4

Molten Globules and Other Compact Denatured States

As mentioned previously, Levinthal (1969) postulated the existence of folding pathways in 1968. In an effort to find the specific intermediates along the folding pathway, folding studies were performed on several globular proteins (Wong and Tanford, 1973; Kuwajima et al., 1976; Robson and Pain, 1976; Nozaka et al., 1978). Equilibrium intermediates were reported but it was not clear whether these are related to the specific folding intermediates. However, the equilibrium intermediates characterized in different proteins were found to be remarkably similar to each other: all of them had native‐like secondary structure and were compact, but lacked a specific tertiary structure. In 1983, Ohgushi and Wada (Ohgushi and Wada, 1983) proposed that the equilibrium intermediates belong to a common physical state of globular proteins, and they termed this the ‘‘molten globule’’ state. After Kuwajima and coworkers (1985) and Ikeguchi and coworkers (1986) had shown that the molten globule state of a‐lactalbumin is identical to its transient folding intermediate, researchers started to study molten globules with renewed interest. Molten globule states could be generated using mild denaturing conditions (low or high pH, moderate concentrations of denaturants, high temperature, and various salts) in about 20–25 different proteins such as a‐lactalbumin, carbonic anhydrase B, b‐lactamase, ribonuclease A, T4 lysozyme, cytochrome c, apomyoglobin, and staphylococcal nuclease (Ptitsyn, 1995).

4.1 The Structure of the Molten Globule The common characteristics of the molten globule state as described by Kuwajima (Arai and Kuwajima, 2000) are (1) the presence of a significant amount of secondary structure, (2) the absence of most of the specific tertiary structure produced by the tight packing of side chains, (3) compactness of the protein molecule with a radius of gyration 10%–30% larger than that of the native state, and (4) the presence of a loosely packed hydrophobic core that increases the hydrophobic surface accessible to solvent. The experimental techniques typically used to detect molten globules are (1) far‐ and near‐UV CD spectra that detect the secondary and tertiary structures of a protein, (2) hydrodynamic methods such as viscosity measurement and molecular sieve chromatography that determine the molecular size of the protein, and (3) hydrophobic dye (typically ANS, 8‐anilino‐naphtalene‐1‐sulfonate) binding experiments that detect the formation of a loose hydrophobic core and estimate the extent of hydrophobic area exposed to the solvent. These techniques, however, only provide information about the average structural properties of the protein molecule. More advanced techniques that provide a more detailed picture about the molten globule were not available until the 1990s. Therefore, the exact nature of molten globule structure had been a matter of some debate. > Figure 10-2 shows an illustration of two different models for the molten globule. The traditional view of the molten globule (Shakhnovich and Finkelstein, 1989; Ptitsyn, 1992) assumes that the backbone of the polypeptide chain essentially has a (fluctuating) native‐like fold and the disorder is in the side chains. However, this view was criticized on the basis of thermodynamic arguments and energy landscape theory (Dill et al., 1995; Privalov, 1996). Privalov (1996) argued that molten globule states are either misfolded structures or states where one portion of the structure is already folded and another one is still unfolded. Dill and coworkers (1995) stated on the basis of simulations of simplified models that backbone and side‐chain degrees of freedom are strongly coupled, therefore a state where the backbone is ordered and the side chains are disordered is unlikely. The new experimental techniques developed in the 1990s, such as stopped‐flow circular dichroism, pulsed hydrogen exchange, X‐ray scattering, and mutational approaches, have allowed characterization of the molten globule states in great detail (Hughson et al., 1990; Dobson, 1994; Carlsson and Jonsson, 1995; Carra and Privalov, 1996; Dabora et al., 1996; Dyson and Wright, 1996; Kataoka and Goto, 1996; Kuwajima, 1996; Song et al., 1998; Chakraborty et al., 2001; Demarest et al., 2001; Ramboarina and

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. Figure 10-2 Illustration of different models for the structure of the molten globule. The backbone of the protein is represented by a thick line and side chains are shown as pieces of various shapes hanging from the backbone. (a) The native state of the protein, with all the buried side chains fitting closely together like the pieces of a jigsaw puzzle. (b) The side chain molten globule model: the fold of the backbone is native‐like but the side chains are only loosely packed. (c) One‐half of the molecule is fully folded, with the specific side chain interactions present, while the other half is completely unfolded

Redfield, 2003; Redfield, 2004). These studies confirmed the view that the molten globule state is indeed more heterogeneous than previously thought: one part of the structure is more organized and native‐like while other portions are less organized. However, there is a great deal of variety among different proteins in the exact structure of the molten globule state.

4.2 The Role of the Molten Globule in Folding The study of molten globules was strongly motivated by the idea that the molten globule state could be identical to the transient intermediate during folding. This notion was supported by kinetic circular dichroism measurements on a‐lactalbumin (Ikeguchi et al., 1986). However, later theoretical studies

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suggested that the experimentally observed folding intermediates may not be productive but kinetically trapped, misfolded species (Dill and Chan, 1997). Careful kinetic measurements during the refolding of several proteins including interleukin‐1b, staphylococcal nuclease, and apomyoglobin (Heidary et al., 1997; Walkenhorst et al., 1997; Jamin and Baldwin, 1998; Maki et al., 1999) have provided firm evidence that, contrary to the theoretical predictions, the molten globule state is the productive on‐pathway folding intermediate in most cases. An attempt to resolve the apparent contradiction between theory and experiment introduced the hierarchical folding model (Arai and Kuwajima, 2000) where the folding of a protein occurs in two stages: (1) formation of the molten globule state from the fully folded state and (2) formation of the native state from the molten globule state.

5

Simulations of Protein Folding and Unfolding

Computational models and simulations have greatly advanced our understanding of protein folding. The information from such studies is complementary to experiments. In fact, there is a synergy between theory and experiment: theory provides testable models and experiments provide the means to test and validate the models. The outcome from this combination is a much richer view of the system in question than what either approach could provide alone. In particular, simulations can help identify or predict transition and intermediate states along the folding pathway, provide predictions of the rate of folding and in some cases, predict the final, folded structure. Simulating protein folding presents a significant challenge. Small proteins typically fold in the several microseconds to seconds timescale; detailed atomistic simulations, however, are currently limited to the nanosecond to microsecond regime. Therefore, simulation of folding requires either simplified models or special sampling methods, both of which introduce new approximations.

5.1 All‐Atom Models The most straightforward approach to simulating protein folding and unfolding is to use an all‐atom model with a force field like AMBER or CHARMM and apply traditional molecular dynamics simulation. These force fields describe the energies of the deformations of covalent bonds as well as van der Waals interactions, charge–charge interactions, hydrogen bonds, and so on. Traditional molecular dynamics numerically solves Newton’s equations of motion by calculating the forces acting on atoms and computing accelerations, velocities, and atomic displacements. Temperature is assigned to the system by assigning appropriate velocities to the atoms. To simulate unfolding, the simplest method of increasing sampling is to increase the temperature of the simulation to 498 K or more. At these temperatures, the native structure of the protein is usually lost within a few nanoseconds. This technique has been applied to numerous proteins, examples of which include bovine pancreatic trypsin inhibitor (Kazmirski and Daggett, 1998a), lysozyme (Kazmirski and Daggett, 1998b), myoglobin (Tirado‐Rives and Jorgensen, 1993), barnase (Wong et al., 2000), ubiquitin (Alonso and Daggett, 1998), the SH3 domain (Tsai et al., 1999), etc. Features of the unfolding process such as the transition‐state ensemble or the unfolded ensemble have shown remarkable agreement with experimental results (Day and Daggett, 2003). A direct simulation of folding is a much harder problem. The longest continuous all‐atom molecular dynamics simulation so far is still the 1 ms simulation of villin headpiece subdomain performed by Duan and Kollman in 1998 (Duan and Kollman, 1998). In this simulation, the chain sampled a large number of conformations after initial collapse, and near‐native structures appeared but the true native conformation was not reached. Although processor speeds have dramatically increased since 1998, computational power is still insufficient to allow a meaningful all‐atom simulation of the entire folding process. Even if the native state could be reached in a single trajectory, multiple simulations would have to be performed to construct a believable folding pathway. An interesting alternative is the massively distributed method employed in the ‘‘Folding at Home’’ project (Pande et al., 2003). A large number of computers run the simulation in parallel. As soon as a

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transition is detected (as a momentary surge in the heat capacity) in one of the simulations, all computers receive a copy of the posttransition conformation and the simulation is continued until the native conformation is reached. Although this approach has been criticized as being flawed (Fersht and Daggett, 2002), it has been successfully applied to fold several small, fast‐folding proteins (Snow et al., 2004; Sorin and Pande, 2005).

5.2 Simplified Models In simplified (coarse‐grained) models (Dokholyan, 2006), effective particles (beads) represent amino acids or groups of atoms. An empirical potential function, usually derived from protein structures, is used to describe the interaction between these beads. The shape of this potential is often very simple, such as a square‐well function. In many simplified models, the positions of the beads are restricted to points on a lattice. Perhaps the most minimal model is the one where there are only two types of amino acids: hydrophobic and polar, and the chain is restricted to a two‐dimensional lattice (Dill et al., 1995). Smaller on‐lattice model proteins allow an exhaustive enumeration of all possible states of the given system. This approach allows a complete thermodynamic description of the phase space and has greatly enhanced our understanding of protein folding. The funnel view and the concept of energy landscapes (see > Sect. 10-6) arose directly from the exhaustive sampling allowed by these minimal models (Bryngelson et al., 1995). In the case of larger, more complex simplified models, exhaustive enumeration of states is not possible. Monte Carlo is a common choice for simulating simplified models. In Monte Carlo simulation, small moves are generated randomly and accepted or rejected based on the energy of the new conformation. This is often performed in the framework of advanced sampling schemes such as Replica Exchange Monte Carlo, where several replicas of the system are simulated at various temperatures (Kihara et al., 2001; Pokarowski et al., 2003). A more recent simulation approach, termed discrete (or discontinuous) molecular dynamics (DMD) (Smith and Hall, 2001; Ding and Dokholyan, 2005), extends the accessible simulation time by using long integration time steps with approximated energy functions. Simple models like this start showing remarkable success. Recently, Trp‐cage, a 20‐residue miniprotein has been folded to a conformation very close to the experimental structure (Ding et al., 2005a). It is believed that this technique will be applicable to larger proteins.

5.3 Multiscale Modeling Approaches using simplified, coarse‐grained models can be combined with fine‐grained, all‐atom simulations in what is called multiscale molecular modeling. Bradley and coworkers (2005) reported high‐ resolution structure prediction for proteins up to 85 residues using a multiscale approach that sampled low‐ and high‐resolution conformations. DMD simulations combined with all‐atom, traditional molecular dynamics have been used to simulate the formation of a b helix (Khare et al., 2005) and to identify the transition state of the SH3 domain (Ding et al., 2005b). Although it remains to be seen whether the force fields used are transferable to larger proteins, multiscale modeling has the potential to break the 1 ms barrier of direct folding simulations.

6

Free‐Energy Landscapes of Proteins

Energy landscapes are mathematical devices that help us understand the microscopic behavior of a molecular system (Bryngelson et al., 1995). An energy landscape of a system with n degrees of freedom is an energy function F(x) ¼ F(x1, x2,. . ., xn) where x1,. . ., xn are variables specifying the microscopic state of the system (Dill, 1999). In the case of a protein, x1, x2,. . ., xn can be, for instance, all the dihedral angles of the chain, thus specifying a single conformation of the protein. F(x) then is usually defined as the free

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energy of the protein in the given conformation, where the entropic part of the free energy comes from all possible solvent configurations. Thus, F(x) is the free energy of a microstate, not a macrostate, because it does not include the chain conformational entropy. The stable conformation of the protein can be found by determining the set of values x1, x2,. . ., xn (i.e., the conformation) that gives the minimum value of the free‐ energy function. Although energy landscapes are, by definition, high‐dimensional surfaces, they are often pictured as a surface in three dimensions. In these pictures, the vertical axis represents the free energy and the horizontal axes represent the conformational degrees of freedom of the polypeptide chain. Random heteropolymers, such as a random sequence of amino acids, have a very rugged energy landscape with many local minima (Plotkin et al., 1996). Systems like this easily get trapped in one of the local minima and usually do not have a well‐defined, single, stable conformation. Real proteins, however, are not random sequences; evolution has optimized their sequences so that they quickly and efficiently fold into a well‐defined three‐dimensional conformation (Onuchic and Wolynes, 2004). In a real protein, most of the interactions that can form between parts of the chain are mutually supportive and cooperatively lead to a low‐energy structure which is therefore ‘‘minimally frustrated.’’ This ‘‘principle of minimal frustration’’ (Bryngelson and Wolynes, 1987), gleaned from simplified models of proteins and the theory of spin glasses, led to the realization that the energy landscape of a real protein should be shaped like a funnel (> Figure 10-3).

. Figure 10-3 Schematic representation of a funnel‐shaped energy landscape. The width of the funnel represents the conformational freedom of the chain. The vertical axis represents the free energy; as free energy decreases, the nativeness of the chain increases. Denatured (unfolded) states are at the top of the funnel while the native state is the global minimum. There is some ruggedness in the energy landscape near the native state

A funnel‐shaped energy landscape means that the free energy of a structure depends on how close it is to the native state: the closer it is to the native structure the lower its free energy. Also, the top of the funnel, representing the nonnative states, is wide (the conformational entropy is high), and it narrows as one gets closer to the bottom: near‐native states represent more compact conformations, and therefore have low conformational entropy. The fact that the energy landscape of a real protein is essentially funnel‐shaped has several important consequences (Wolynes, 2005). First, the native structure should be robust against mutations (Nelson and Onuchic, 1998). A point mutation represents a small perturbation to the energy landscape; therefore, the basic shape of the funnel and the location of the global minimum cannot change much: the mutant protein will fold into essentially the same structure as the wild‐type protein. This structural robustness underlies the most commonly used method of predicting structures: homology modeling (Zhang and Skolnick, 2004).

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If sequence analysis shows two sequences to be evolutionarily related, one can be almost certain that the two structures are essentially the same. Another important consequence of the funnel shape is that the folding rate and folding mechanism of a protein is largely determined by its native‐state topology (Baker, 2000). Assuming that the energy landscape is a perfect funnel, its shape is completely determined by the topology of the native state, and this shape completely determines the folding mechanism. Of course, the energy landscape of a real protein is never a perfect funnel: it is always somewhat rugged, and the exact layout of the small heaps and valleys on the landscape will influence the details of the folding mechanism. Still, for many proteins, the landscape is close enough to a perfect funnel to allow the development of methods that can successfully predict various features of the folding mechanism from just the native structure as input (Alm and Baker, 1999; Galzitskaya and Finkelstein, 1999; Munoz and Eaton, 1999). The folding rate can be predicted from just the contact order (the average sequence separation between residues that make contacts in the native structure). Using a perfectly smooth, funnel‐shaped energy function based on the Go model (Go, 1983) (where only native contacts contribute to the free energy), transition‐state structures, folding nuclei, and F values have been predicted for several small proteins such as CheY, CI‐2, barnase (Galzitskaya and Finkelstein, 1999), l‐ repressor, and SH3 domain (Alm and Baker, 1999), with surprisingly good agreement with experiments. The concept of energy landscapes also helps us understand some of the more complex protein‐folding scenarios. The folding of many proteins involves slow steps, bottlenecks, and multiple, kinetically distinguishable stages (Wolynes et al., 1995). These phenomena can be rationalized by noticing that energy landscapes are often not perfect, smooth funnels but are rugged and bumpy (Dill, 1999). Slow steps in the folding process can arise from climbing an uphill slope after being trapped in local minima. This scenario may appropriately describe the folding of b‐lactoglobulin, a predominantly b‐sheet protein, which passes through a helical phase as it folds (Hamada et al., 1996). Because the landscape view is a microscopic view of the folding process, each molecule may follow a different route on the energy landscape and may encounter different obstacles in its way. For example, in the folding of hen egg white lysozyme, a subpopulation of the molecules folds fast while another subpopulation folds quite slowly (Radford et al., 1992). In the landscape view, it is easy to see how molecules starting from one region of the top of the funnel may ‘‘ski down’’ unhindered while molecules starting from another region may get trapped behind a mountain range. Bottlenecks do not always involve uphill climbing though, they can be entropic barriers too; in this case, the aimless search for a downhill route on a large, level field will limit the rate of folding (Dill and Chan, 1997). This example shows that bottlenecks are typically ensembles of widely different conformations rather than well‐defined, single conformations (Chan and Dill, 1998). The landscape view of protein folding also gives us a clue to chaperone function: to get a misfolded protein to fold correctly, no specific recognition is needed; it is sufficient just to move it to the top of the funnel where it can restart the downhill search for the folded conformation (Chan and Dill, 1996; Todd et al., 1996).

7

Traps on the Folding Pathway

The folding protein faces several obstacles in translating the information encoded in the amino acid sequence into the three‐dimensional construct of the biologically active structure. The cis–trans isomerization of peptide bonds and the formation of disulfide bridges are slow steps that can form bottlenecks in the protein‐folding reaction (Balbach and Schmid, 2000; Creighton, 2000). Under certain circumstances, the two processes can be linked and the formation of the correct disulfide bonds is facilitated in the presence of peptidyl‐prolyl cis–trans isomerase (Schonbrunner and Schmid, 1992).

7.1 Backbone Isomerization The peptide bond between nonproline amino acids is much more stable in the trans than in the cis conformation (> Figure 10-4a). The difference in stability arises from interactions between the Cai and Hai atoms with the Cai þ 1 and Hai þ 1 atoms, from electrostatic interactions between the Oi and Ci þ 1 atoms

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. Figure 10-4 (a) The cis and trans states of nonprolyl peptide bonds. (b) The cis and trans states of prolyl peptide bonds

and from conformational entropy (Zimmerman and Scheraga, 1976; Wedemeyer et al., 2002). The large difference in the stability of the two isomers keeps the trans form 100 to 1000 times more populated than the cis isomer (Ramachandran and Mitra, 1976; Jorgensen and Gao, 1988; Scherer et al., 1998). Because of the high number of peptide bonds in the protein, the fraction of protein molecules that have cis peptide bonds in the denatured ensemble can be significant. Since isomerization of peptide bonds is a slow process, nonnative isomers of peptide bonds can cause slow phases in protein folding. It has been shown that cis–trans isomerization of nonprolyl peptide bonds can give rise to significant slow folding phases (Eyles, 2001). Nonprolyl cis peptide bonds are energetically unfavorable and are very rare in the native structures of proteins (Stewart et al., 1990). Isomerization of the trans peptide bond accumulated in the denatured state into the native cis conformation of these proteins can become the rate‐limiting step of the folding reaction (Odefey et al., 1995). Proline residues are a much more common source of kinetic complications during folding. The X‐Pro peptide bond (where X can be any amino acid) is the only peptide bond for which the stability of the cis and trans conformations is comparable. The cis–trans isomerization of X‐Pro peptide bonds is a widely recognized rate‐limiting factor, which can induce additional slow phases in protein folding (Brandts et al., 1975; Wedemeyer et al., 2002). The otherwise strong preference of the peptide bond to be in trans state does not apply to the X‐Pro bond, mostly because of the steric symmetry between the Ca and the Cg atoms of proline (> Figure 10-4b). The entropy change accompanying the cis–trans transition and the electrostatic interactions between the Oi and Ci þ 1 atoms are also different in the X‐Pro bonds than in other peptide bonds (Zimmerman and Scheraga, 1976; Wedemeyer et al., 2002). As a result, the trans conformation of X‐Pro bonds is only marginally more stable than the cis conformation in unfolded polypeptides. In the unfolded protein, a mixture containing both conformations is present, with 10%–30% of the X‐Pro bonds in the cis state (Cheng and Bovey, 1977; Grathwohl and Wuthrich, 1981). In the native conformation of a protein, intraprotein interactions will stabilize either the cis or the trans isomer for most of the X‐Pro bonds. The molecules that contain incorrect isomers in the unfolded ensemble must undergo isomerization during the folding reaction (Kiefhaber et al., 1990a, b; Texter et al., 1992). Consequently, the presence of proline residues leads to a number of proline isomerization reactions that must occur before folding can complete. Both experimental and theoretical findings show that there is a high energy barrier for isomerization (16–20 kcal/mol in model compounds) of X‐Pro bonds (Balbach and Schmid, 2000; Kang and Choi, 2004). This results in characteristic times of 10–1000 s for the

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conformational flipping at room temperature (Brandts et al., 1975). Mutagenesis studies have indeed shown that specific proline residues can often be assigned to slow recovery phases from misfolded states (Evans et al., 1987; Kelley and Richards, 1987; Wood et al., 1988; Herning et al., 1991; Kiefhaber et al., 1992; Wu and Matthews, 2002; Street et al., 2005). The high energy barrier for isomerization reflects a partial double‐bond character of the X‐Pro bond (Balbach and Schmid, 2000). In vivo, enzymes (e.g., peptidyl‐prolyl cis–trans isomerases) facilitate folding to the native structure by lowering the energy barrier for isomerization (Schmid et al., 1993; Stein, 1993; Gothel and Marahiel, 1999; Shaw, 2002). Laser‐induced temperature‐jump measurements indicate that the role of proline residues in protein folding is more complex. Prolines can have opposite effects on the slow and fast steps of the folding kinetics. The presence of proline residues leads not only to additional slow phases, but also modifies the millisecond and sub‐millisecond dynamics of the protein. The X‐Pro bonds do not isomerize on the millisecond timescale. This increased backbone rigidity can speed up the fast folding steps of the ensembles that contain the proline in the native‐like isomerization state (Osvath and Gruebele, 2003).

7.2 Formation of Disulfide Bridges Formation of the correct native‐like disulfide bridges can also hinder protein folding. Disulfide bridges between pairs of cysteines are part of the native structure for many proteins. The formation of disulfide bridges is a prerequisite of the proper folding and biological function in these proteins. Disulfide bonds increase the thermodynamic stability of the native structure by establishing conformational constraints within the protein (Creighton, 2000). It has been shown that several proteins start to fold during synthesis and disulfide bond formation begins in the emerging chain (Bergman and Kuehl, 1979; Peters and Davidson, 1982; Braakman et al., 1991; Braakman et al., 1992). Folding and disulfide bridge formation is completed after the end of the translation (Wedemeyer et al., 2002). Depending on the number of the cysteine residues in a polypeptide sequence and on the native structure, misfolding can occur owing to errors in disulfide pairing. Disulfides formed randomly in early folding intermediates may cause kinetic complications during the later folding steps (Creighton, 1979; Konishi et al., 1982). Repairing of the errors is part of the folding process and occurs in a trial and error manner (Chatrenet and Chang, 1992; Schwaller et al., 2003). Since disulfide bonds constrain the free movement of the polypeptide chain, the formation of native disulfide pairs can also prevent correct folding. It has been shown that native‐like disulfide bridges formed too early during folding must be broken up to allow the folding process to continue, and they can reform at a later stage (Creighton and Goldenberg, 1984). Formation of disulfide bonds requires the presence of an oxidizing agent or a disulfide reagent such as glutathione or dithiothreitol. When a disulfide reagent is present, disulfide bond formation occurs by a thiol–disulfide exchange reaction. This is a two‐electron redox reaction in which a disulfide reagent takes one electron from each cysteine thiolate, and a disulfide bond is formed (> Figure 10-5) (Wedemeyer et al., 2000). The first step is the result of a nucleophilic attack by a cysteine thiolate on the disulfide reagent. The rate of this reaction is determined by the reactivity of the cysteine thiolate, the presence and nature of the disulfide reagent, pH, temperature, ionic strength, and cosolvents. In the second step, the mixed disulfide is broken up by another nucleophilic attack by the second cysteine thiolate. As a result, the external thiol is replaced by a protein thiolate and an intraprotein disulfide bond is formed. The prerequisite of the second step is a conformational change in the protein that brings the two cysteine residues together, thus the redox reaction is coupled to the folding of the protein (Creighton, 1997; Bulaj, 2005). The disulfide pairing during in vivo folding is helped by protein disulfide isomerases (Bulleid and Freedman, 1988; Freedman, 1989; Ellgaard and Ruddock, 2005). These enzymes correct the disulfide pairing mistakes by catalyzing the reshuffling of disulfide bridges so that the native pairing can emerge (Gilbert, 1997). The process leads to a biologically active structure, which is resistant to further rearrangement (Walker and Gilbert, 1997).

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. Figure 10-5 Scheme of the redox reaction that leads to the formation of disulfide bonds in proteins

8

Transition States on the Folding Pathway

To gain insight into the folding mechanism, it is essential to characterize the energy landscape of the protein. This includes the determination of the structures and energies of the protein in the traps and the transition states of the landscape (Gruebele, 2002). The local minima of the energy landscape act as traps that are responsible for the buildup of kinetic intermediate states. Since the protein accumulates in measurable quantities in the intermediate states, the intermediate structures can be experimentally studied (Roder et al., 2000). Transition states are usually not populated in detectable quantities. The only way to gain information about the structural and energetic properties of the transition states is through kinetic studies of the folding reaction (Daggett and Fersht, 2000). The kinetics of the transition between different states is determined by the escape rates from local minima of the landscape. In simple cases, protein folding can be described as a diffusive process over a barrier determined by the energy landscape. For barriers that are larger than the thermal energy kBT, the folding rate predicted by transition‐state theory can be calculated by a Kramer‐like equation (Onuchic et al., 1996) . k ¼ n exp DG { kB T : Here k denotes the rate of the conformational transition, n is a coefficient that depends on a number of things including the shape of the barrier and solvent viscosity, DG{ is the Gibbs free‐energy height of the transition state, kB is Boltzmann’s constant, and T is the absolute temperature. Transition‐state theory thus allows us to determine the Gibbs free energy of the transition state using simple kinetic measurements. Gaining information about the structure of the transition state is a more intricate problem. To date, the only way to learn about the structure of the transition state is F‐value analysis, a method that uses site‐directed mutagenesis to map out the residue–residue contacts present in the transition state (Matthews and Hurle, 1987; Fersht et al., 1992). The calculated F‐value compares the change in the folding rate and the change in the stability of the protein caused by a specific mutation (Clementi et al., 2000) F ¼ ðRT lnðkmut =kwt ÞÞ DGmut : DGwt Here R is the gas constant, T is the absolute temperature, and kmut and kwt are the folding rates of the mutant and wild‐type protein, respectively. DGmut and DGwt represent the stabilities of the mutant and of the wild‐type protein, i.e., the Gibbs free‐energy difference between the folded and the unfolded state. If the unfolded state is assumed to be a randomly fluctuating chain, its free energy can be taken as the solvation free energy. However, it has been shown that denatured proteins can have some residual structure (Smith et al., 1996; McCarney et al., 2005), and a determination of their free energies becomes more complicated. For the purposes of F‐value analysis, however, we can just use the Gibbs free energy of the unfolded state as a reference, i.e., define it as zero. The expression for F can be simplified if the following two conditions are met: (1) The folding rate can be calculated from the activation energy using a Kramer‐like exponential dependence, thus folding can be treated as a simple cross‐barrier diffusive reaction and (2) The folding

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mechanism is not altered significantly by the mutation, thus the parameter n reflecting the barrier shape and the configurational diffusion of the protein is insensitive to the mutation. If the above assumptions hold, the new expression for F is: . { { DGmut ; DGwt DGwt F ¼ DGmut where DG{mut and DG{wt denote the activation Gibbs free energies for the mutant and the wild‐type protein, respectively. This way, the F‐value compares the Gibbs free‐energy change introduced by the mutation in the transition state and in the native state (> Figure 10-6).

. Figure 10-6 Schematic representation of the Gibbs free energies important in the F‐value analysis

In order to be able to extract structural information from F‐value analysis, two more conditions have to be met (Fersht et al., 1992; Clementi et al., 2000): 1. The folding pathway is not altered significantly by the mutation, thus the intermediate and transition states are the same for the mutant and the wild‐type protein. 2. The folded region of the transition state has a native‐like structure. The above conditions were found to be true for several proteins, and F‐value analysis was used to characterize transition states and short‐lived intermediates of several folding pathways (Matouschek et al., 1990; Serrano et al., 1992; Grantcharova et al., 1998; Villegas et al., 1998; Fulton et al., 1999; Goldenberg, 1999; Raschke et al., 1999; Crane et al., 2000; Bulaj and Goldenberg, 2001; Capaldi et al., 2002; Paci et al., 2003; Hubner et al., 2004; Lindorff‐Larsen et al., 2004; Hubner et al., 2005). Usually, a detailed analysis of the mutation is necessary to identify the contacts disrupted in the mutant structure. The use of multiple mutations of the same residue can increase the accuracy of the prediction of the transition‐state structure (Matouschek et al., 1995). F‐values are normally between 0 and 1, but negative F‐values corresponding to mutations that speed up the folding kinetics have also been observed (Yang and Gruebele, 2003; Yang and Gruebele, 2004b). The F‐value is a valuable tool in detecting native contacts in transition‐state structures, but in general, it is not proportional to the extent or strength of the native contact appearing in the transition state. This means that structural information can unambiguously be assigned only to F ¼ 0 and F ¼ 1 (Fersht and Sato, 2004). A F‐value close to 1 indicates that the free‐energy change introduced by the mutation is almost identical for the transition state and the native state. This implies that the mutated residue already forms its native

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contacts in the transition state. A F‐value close to 0 indicates that the height of the Gibbs free‐energy barrier of the transition state was not altered by the mutation. In this case, the mutated residue does not form native contacts in the transition state, and the environment of the mutated residue is probably denatured‐ like (Fersht and Sato, 2004). It has been shown that for deletions of small hydrophobic residues, the F‐value is roughly proportional to the extent of the native contact formation. Therefore, these residues are preferred as targets for mutations in F‐value analysis (Fersht and Sato, 2004).

9

Folding of Multidomain and Multi‐Subunit Proteins

9.1 Multidomain Proteins A domain is a part of the polypeptide chain that forms a compact globular substructure with more interactions within itself than with other parts of the chain (Janin and Wodak, 1983). Most proteins longer than about 200 to 250 residues consist of several domains. When investigating the role of domains in folding, the first question to answer is whether a domain can fold by itself. Isolated domains can be produced by limited proteolysis or genetic engineering, and their folding can be studied by the same experimental methods as used for the whole proteins. Autonomous folding was demonstrated for the domains of several multidomain proteins including tryptophan synthase, b‐lactamase, aspartokinase– homoserine dehydrogenase, plasminogen, phosphoglycerate kinase (Jaenicke, 1987), and several other domains (Sharma et al., 1990; Herold et al., 1991; Shoelson et al., 1993; Williams and Shoelson, 1993; Jecht et al., 1994). In many cases, it was shown that the stability of the domains in isolation is close to the stability when the other domains are also present (Garel and Dautry‐Varsat, 1980; Muller and Garel, 1984; Novokhatny et al., 1984; Jaenicke, 1987; Tsunenaga et al., 1987; Rudolph et al., 1990; Missiakas et al., 1992). Interestingly, the rate of folding of isolated domains was found to be greater or the same as that of the same domains integrated within the intact protein (Teale and Benjamin, 1977; Dautry‐Varsat and Garel, 1981; Blond and Goldberg, 1986; Tsunenaga et al., 1987; Missiakas et al., 1992). This suggests that the presence of the rest of the chain slows down the folding of a domain, e.g., by forming unfavorable interactions with it. The folding kinetics of multidomain proteins is usually complex, showing an initial rapid phase characterized by large changes in several physical parameters (fluorescence, UV absorption, circular dichroism, etc.), followed by a second, slower phase by much smaller changes in the physical parameters (Jaenicke, 1987; Jaenicke, 1999). The intermediate that accumulates after the initial rapid phase appears largely folded but lacks some properties of the native structure: it is more labile to proteolysis, is not recognized by some antinative antibodies, and lacks catalytic activity. These properties only appear after the second, slower stage of folding. The findings suggest that the individual domains fold in the rapid phase of the folding process, and the second, slow step is the pairing or association of the already folded domains. This is also supported by the fact that in several cases the rate of folding was found to be inversely proportional to solvent viscosity, implying that movement of large, globular species is involved in the rate‐limiting step of the folding process (Vaucheret et al., 1987; Chrunyk and Matthews, 1990). Also, in tryptophan synthase, single mutations in each domain were found to decrease the folding rate while the double mutant was able to fold at the same rate as the wild‐type protein, suggesting that the rate‐limiting step of folding involves the formation of the interface between the two domains (Tsuji et al., 1993). Thus, the folding process can be described by the following general scheme Fast domain folding

Slow domain pairing

Unfolded ! Intermediate with folded but unpaired domains ! Native The existence of an intermediate with folded but unpaired domains has important consequences. If the protein concentration is sufficiently high, the domain pairing step may occur between domains from two identical molecules, leading to the formation of a dimer instead of a monomer. The domains make the same interactions in the dimer as in the monomer, but the interactions are formed intermolecularly rather than

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. Figure 10-7 The folding of a hypothetical two‐domain protein. The unfolded chain folds into two domains that are not yet paired. The fate of this intermediate depends on the protein concentration and the solvent conditions: it may either form the native state; it may associate with another chain to form a domain‐swapped dimer; or several chains may aggregate or form a fibril

intramolecularly. Higher‐order oligomers may also form and the process may lead to fibril formation or aggregation (see > Figure 10-7). The exchange of domains between identical molecules is actually a special case of a range of phenomena termed ‘‘domain swapping’’ (Bennett et al., 1994). The term describes situations where two or more proteins exchange part of their structure (not necessarily whole domains) to form intertwined oligomers (Rousseau et al., 2003). Domain swapping has been implicated in amyloid fibril formation, although not all models of amyloid formation involve domain swapping (Nelson and Eisenberg, 2006).

9.2 Multi‐Subunit Proteins Almost all large proteins are formed by the association of subunits. There are several evolutionary advantages to forming multimers: regulation of catalytic activity through cooperativity between the subunits, generation of new functions and enzymatic activities, formation of large structures, increasing

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protein stability, and facilitating the folding process. Subunit–subunit interfaces are very diverse regarding the nature and distribution of inter‐subunit interactions. About one‐third of the interfaces have a large and contiguous hydrophobic patch surrounded by a ring of inter‐subunit polar interactions; the remaining two thirds show a mixture of small hydrophobic patches, polar interactions, and water molecules scattered over the interface area. There are several experimental techniques to study the folding and association of multi‐subunit proteins. Depending on the strength of the binding between the subunits, the experimentalist can attempt to produce folded but dissociated subunits under equilibrium conditions, using techniques such as dilution, cold dissociation, chemical modification, ligand‐induced dissociation, mildly denaturing conditions, or elevated pressure (Jaenicke and Lilie, 2000). If unfolded monomers cannot be obtained, folding and association should be studied as coupled processes. Several methods can be used to monitor the association state of the protein after starting reassociation of the (folded or unfolded) subunits. By rapid chemical cross‐linking, snapshots can be taken during the reassociation process and investigated by gel electrophoresis. Other methods include hybridization with isoenzymes or modified subunits and measurement of relative reactivation. There is also a wide range of biophysical methods to monitor and/ or measure the thermodynamic parameters of protein–protein interactions (Lakey and Raggett, 1998), including surface plasmon resonance, isothermal titration calorimetry, fluorescence energy transfer, mass spectrometry, light scattering, and high‐pressure liquid chromatography. After collecting the time‐ dependent data, a reaction equation can be fit and a reaction scheme established. The assembly of multi‐subunit proteins can usually be described as a series of unimolecular (isomerization) and bimolecular (association) steps 4Mu ! 4M ! 2D0 ! 2D ! T0 ! T; where M represents a monomeric, D a dimeric, and T a tetrameric state, and the subscript ‘‘u’’ represents the unfolded state. Are folding and association separate events, or are they more or less coupled? The traditional view is that monomers assume a near‐native conformation before binding to their partners. In recent years, however, several proteins have been found that are intrinsically unstructured as monomers and only fold upon binding to DNA, RNA, a membrane, or another protein (Dyson and Wright, 2002). Homodimers whose monomers only fully fold upon dimerization include troponin C site III (Monera et al., 1992), Arc repressor (Robinson and Sauer, 1996), FIS (factor for inversion stimulation) (Hobart et al., 2002), Trp repressor (Gloss et al., 2001), and the dimeric form of p53 (Mateu et al., 1999). The HIV gp41 protein is a homotrimer with intrinsically unfolded monomers (Marti et al., 2004). It has been shown that the binding mechanism (whether monomer folding is coupled to binding or there are folded monomeric intermediates) is determined by the native topology, especially the number of inter‐ and intramolecular contacts and the hydrophobicity of the interface (Levy et al., 2004). Flexibility of the chain seems to play a major role in the binding mechanism (Levy et al., 2005); one important manifestation of this is the fly‐casting effect (Shoemaker et al., 2000): a relatively unstructured chain can have a greater ‘‘capture radius’’ to ‘‘catch’’ its binding partner before fully folding.

10

Protein Folding in the Cell

Under carefully chosen in vitro conditions, small single‐domain proteins fold in a cooperative and reversible manner. Usually, the experiment is carried out in dilute solutions at low temperatures where the folding reaction is not complicated by off‐pathway reactions such as aggregation. Contrarily, the interior of a cell is a highly crowded environment with an estimated protein concentration of 300 mg/ml (Zimmerman and Trach, 1991), and in extreme cases, such as in a thermophilic archaeon, the temperature can exceed 80 C. Another difference is that in vivo the folding process is not separated from the relatively slow synthesis of the polypeptide chain (Creighton, 1990; Jaenicke, 1991). Folding cannot complete before a folding unit is completely synthesized. Nascent chains emerging from the ribosome should avoid formation of misfolded intermediates and aggregation. To ensure efficient folding, cells have evolved a large and

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diverse group of proteins that assist the formation of the native structure. These proteins form the complex machinery of ‘‘molecular chaperones,’’ whose function is to guide other proteins to their proper folding and unfolding routes and help the assembly or disassembly of macromolecular structures, without becoming permanent components of these structures. Some chaperones are also stress or heat‐shock proteins because the need for chaperone function increases under conditions of stress that cause proteins to unfold or misassemble (Ellis, 2005). Although chaperone systems in eukaryotic cells are more complex than in prokaryotes, several homologous classes of chaperones have been identified in these different cell types. Two major classes of chaperones, the Hsp70s and the cylindrical chaperonin complexes, protect nonnative polypeptide chains from misfolding and aggregation in the cytosol of prokaryotes and eukaryotes, and inside the chloroplasts and mitochondria (Ostermann et al., 1989; Hartl, 1996), in an ATP‐dependent manner. The two classes exhibit distinct structural and functional properties and show entirely different mechanisms of action. Other protein classes, such as the Hsp40s, nucleotide exchange factors, or ADP‐destabilizing factors, act as cofactors to the chaperones. Here, we briefly summarize what is known about the major chaperone classes.

10.1 The Hsp70 Family Hsp70s are distributed in all types of cells and in all cellular compartments. The Hsp70 family is a highly conserved protein family consisting of numerous homologs with distinct cellular functions (Frydman, 2001). These chaperones have a molecular mass of approximately 70 kDa. The molecules consist of two domains, the 44 kDa N‐terminal domain, which mediates ATP binding (Flaherty et al., 1990) and the small C‐terminal domain, which binds the protein substrate (Zhu et al., 1996). Substrate binding and release is modulated by ATP binding and hydrolysis. When ATP is bound, substrate binding and release occur rapidly, while with ADP bound, both substrate binding and release are slow. The function of Hsp70s is tuned by cofactors modulating substrate and nucleotide binding. Members of the Hsp70 family are DnaK in bacteria, Ssa and Ssb in yeast, and Hsc70 and Hsp70 in mammals. Hsp70s have a substrate‐binding cleft, which recognizes extended stretches of polypeptide chains rich in hydrophobic residues (Flynn et al., 1991; Rudiger et al., 1997), such as segments of partially unfolded proteins in nonnative conformations. Bound substrates are protected from aggregation; however, folding is obviously not possible in the bound state. Repeated binding and release might keep substrate proteins in extended, monomeric conformation giving them the chance to assume their native structure. However, many of the Hsp70‐bound substrates are transferred to the real ‘‘folding machine,’’ the chaperonin system. Hsp40s are cofactors that stimulate ATP hydrolysis by Hsp70s. Hsp40s are also capable of binding substrates and can pass the bound substrates on to a Hsp70 molecule. The molecule usually consists of an N‐terminal J‐domain, which is responsible for Hsp70‐binding and a C‐terminal chaperone domain containing hydrophobic patches for substrate binding (Sha et al., 2000). Representatives of the family are DnaJ in E. coli, Ydj1 and Sis1 (ribosome associated) in yeast, and Hdj1–2 and Hsp40 in mammals. Several eukaryotic homologs contain only a J‐domain, which may have a role in the localization of Hsp70s. Nucleotide exchange factors promote the release of ADP from members of the Hsp70 family. This group includes GrpE, a 23‐kDa protein in E.coli, and its homologs in mitochondria and chloroplasts, as well as several 60‐kDa proteins including Sti1 in yeast and Hop in mammals. In mammals, the Bag1 protein (Hohfeld and Jentsch, 1997; Takayama et al., 1997) also promotes the release of ADP from Hsp70 and the subsequent substrate release. These are modular proteins with domains that might have a role in connecting different chaperone systems. Hop may link Hsp70 to the Hsp90 system (Gross and Hessefort, 1996), and Bag1 contains an ubiquitin homology domain, suggesting a possible direction of Hsp70‐bound substrates to the S26 proteasome system (Luders et al., 2000; Terada and Mori, 2000). ADP stabilizing factors, such as the 48‐kDa Hip protein in mammals, bind Hsp70 and stabilize the ADP–Hsp70 complex.

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Small chaperones exhibiting Hsp70‐like activity have a function similar to that of Hsp70, but unlike Hsp70, their function is independent from nucleotide binding. The trigger factor (TF) in E. coli has an overlapping function with DnaK. Additionally, it binds to the ribosome and displays prolyl‐isomerase activity (Hesterkamp et al., 1996). Hsp70 can be functionally substituted by the ubiquitous prefoldin (GimC) chaperone. Prefoldin is a heterohexamer that can interact with nascent polypeptides in vitro. The crystal structure of prefoldin reveals a unique quaternary structure forming a novel class of chaperones (Siegert et al., 2000).

10.2 The Folding Cage of Chaperonins (Hsp60 Family) Chaperonins are barrel‐like multi‐subunit complexes that primarily promote ATP‐dependent protein folding (Farr et al., 2000; Brinker et al., 2001). The unique mechanism of chaperonins involves the capture and isolation of substrate polypeptide chains inside the chamber of the complex. In the case of group I chaperonins such as GroEL in E. coli, the mitochondrial Hsp60, and the RuBisCo‐ binding subunit (RBP) in plants, the system requires a Hsp10‐type co‐chaperonin that acts as a ‘‘cap’’ or ‘‘lid.’’ The GroEL system with GroES as co‐chaperonin is capable of correctly folding proteins of sizes up to 60–70 kDa and with multiple domains (Houry et al., 1999). GroEL is a homooligomer of 14 subunits (see > Figure 10-8). These subunits consist of an apical, an intermediate, and an equatorial domain and are arranged in two stacked rings forming two chambers. The apical domains in the open conformation

. Figure 10-8 (a) The structure of the GroEL complex with the GroES lid on (closed state). (b) The structure of the GroEL complex in the open state (no GroES lid present). One subunit of both GroEL and GroES is shown in a darker color

provide hydrophobic side chains that can interact nonspecifically with the exposed hydrophobic surface of the unfolded substrate chain. Subsequent binding of the GroES cap and seven ATP molecules to GroEL triggers a conformational change resulting in an increased volume of the central cavity, a separation of the hydrophobic residues, and an exposure of hydrophilic residues in the apical region (Shtilerman et al., 1999). This may induce partial unfolding of the substrate molecule and its release into the inside of the central cavity of the chaperonin. The isolated environment of the central cavity is ideal for the substrate molecule for folding up into its native structure. Upon completion of ATP hydrolysis, which may take approximately 10 s, binding of seven new ATP molecules to the trans ring triggers the dissociation of the GroES cap and the substrate molecule is released. In the case of incomplete folding, the substrate molecule can be recaptured and the ‘‘annealing’’ and relaxation cycle can be repeated until the molecule correctly folds.

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Group II chaperonins are found in eukaryotes and in archaea and are homologous to group I chaperonins with a sequence identity up to 40%, showing similar double‐ring architecture. The eukaryotic group II chaperonin named TCP‐1 (for tailless complex polypeptide‐1) or CCP (for chaperonin‐containing TCP‐1) is a heterooctamer consisting different subunits of 55–60 kDa (Kubota et al., 1995). Group II chaperonins have a helical protrusion on the apical domain that takes the place of the co‐chaperonin GroES. The crystal structure of the thermosome from Thermoplasma acidophilum reveals that this protrusion can assume different conformations in the open state including a‐helix and b‐sheet, which can increase the plasticity for binding different types of substrates.

10.3 The Hsp90 Chaperone System Hsp90 plays a central role in eukaryotes in the regulation of the components of signal transduction systems such as tyrosine kinases and steroid hormone receptors. Hsp90 is ATP dependent and requires interaction with several cofactors, some of them having chaperone activity. An example is the substrate transfer from Hsp70 to Hsp90, which is facilitated by Hop (Hsp70–Hsp90‐organizing protein). Hop contains binding sites for both chaperones and contributes to the rapid transfer of receptor molecules from Hsp70 to Hsp90. A detailed review on Hsp90 function has been published recently (Pearl and Prodromou, 2006).

10.4 Chaperone‐Assisted Assembly of Cellular Complexes Recent work has shown the role of nuclear chaperones in the assembly of nucleosomes and has led to the discovery of a cytosolic chaperone required for mammalian proteasome assembly, suggesting that besides the folding of individual proteins, the formation of oligomeric complexes may also be assisted by chaperones (Ellis, 2006).

10.5 Cooperation Between the Different Chaperone Systems Newly synthesized polypeptide chains, proteins losing their native state upon stress conditions, and other proteins requiring translocation in the cell may interact with several different chaperone systems. Chaperones having overlapping functions may compete for their substrate molecules as well. Hsp70 binds polypeptides and prevents aggregation. This may be sufficient for the correct folding of some proteins. Others may require further transfer from Hsp70 to the more efficient folding machine: the Hsp60 system. In E. coli, the cooperation between DnaK and GroEL has been proven in vivo (Teter et al., 1999). TF, which may substitute for the function of DnaK, also appears to cooperate with GroEL in substrate binding (Kandror et al., 1997). In eukaryotic cells, Hsp70 and TCP‐1 associate, indicating a close functional relationship (Lewis et al., 1992). As mentioned above, Hsp70 and Hsp90 may also interact in receptor transfer. > Figure 10-9 shows a schematic representation of the general view of de novo protein folding in the cytosol.

10.6 Chaperones and Neurodegenerative Diseases The pathogenesis of several neurodegenerative diseases is associated with protein misfolding, aggregation, and deposition of the protein, which may be manifested in cell degeneration and loss of function of the affected cells or organs. These degenerative disorders include polyglutamine (polyQ) tract diseases, Alzheimer’s and Parkinson’s disease, amyotrophic lateral sclerosis, and Creutzfeldt–Jakob syndrome (Chiti and Dobson, 2006). Chaperones whose function is to prevent protein aggregation in the cell may play a crucial role in the onset or progress of neurodegenerative diseases.

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. Figure 10-9 A schematic representation of the general view of de novo protein folding in the cytosol

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Protein Misfolding and Aggregation

Native states of proteins almost always represent the thermodynamically most stable conformation under physiological conditions (Vendruscolo et al., 2003). All the information regarding the native structure is hidden in the amino acid sequence. However, as we have seen, correct folding is a challenge for proteins in a living cell and only a part of the proteins can assume their native structure spontaneously. In the crowded milieu of the cell, efficient protein folding and transport depend on the presence of a complex machinery of chaperones, chaperonins, and cofactors. The primary mission of this machinery is to prevent the aggregation of nascent polypeptide chains and proteins that unfold upon environmental stress (Frydman, 2001). The failure of a specific protein to adopt or maintain its native functional conformation may result in pathological conditions referred to as ‘‘protein misfolding diseases.’’ Misfolding diseases include a wide range of diseases with different pathological mechanisms. The loss of the normal cellular function because of a reduction in the number of functional protein molecules is responsible for diseases such as cystic fibrosis (Amaral, 2004) and early‐onset emphysema (Lomas and Carrell, 2002). The major group of misfolding diseases, however, is associated with aggregation and deposition of proteins in the human body in the form of organized, fibrillar aggregates, generally termed amyloid fibrils. On one hand, unwanted protein aggregation may be caused by malfunctioning of the cellular protein quality‐control machinery. It may occur when the ubiquitin‐proteasome protein degradation system cannot eliminate misfolded, aggregation prone molecules (Ross and Pickart, 2004; Mandel et al., 2005); when the chaperone machinery performs insufficiently (Lee and Tsai, 2005); if the normal cellular transport route of a protein is damaged; or when inappropriate protease activity produces amyloidogenic protein fragments. On the other hand, aggregation and amyloid formation of a protein may be promoted by an increased expression level or by pathological mutations destabilizing the structure and inducing intermediate amyloidogenic conformations (Chiti and Dobson, 2006).

11.1 Degenerative Diseases Associated with Amyloid Deposition Amyloid deposition is associated with more than 20 human degenerative diseases. We distinguish different groups of diseases by the location of amyloid deposits in the body. Neurodegenerative conditions such as Alzheimer’s, Parkinson’s, and Huntington’s disease as well as spongiform encephalopathies affect the central nervous system. In nonneuropathic localized amyloidoses, protein deposition occurs in a certain type of tissue such as Langerhans’ islands in type II diabetes. In systemic amyloidoses such as AL amyloidosis, which involves the deposition of immunoglobulin light‐chain fragments, protein deposition is not limited to a single tissue. Alzheimer’s and Parkinson’s disease are sporadic and usually develop with aging, which suggests the role of the protein quality‐control machinery in the disease, although hereditary forms are also documented. Other conditions are hereditary, arising from specific mutations, such as lysozyme and fibrinogen amyloidosis. The special property of spongiform encephalopathy is that it can be transmissible in humans and mammals (Chiti and Dobson, 2006). Hemodialysis‐related amyloidosis represents the first amyloid disease that is a complication of a medical therapy (Gejyo et al., 1985). Protein aggregates can accumulate both extracellularly and intracellularly. The term ‘‘intracellular inclusion’’ has been suggested as more appropriate for amyloid‐like aggregates depositing inside the cell (Westermark et al., 2005). However, herein we use the term ‘‘amyloid fibril.’’

11.2 The Structure and Morphology of Amyloid Fibrils A common characteristic feature of amyloid fibrils formed from different proteins is the well‐ordered structure with high b‐structure content. X‐ray fiber diffraction studies have shown that the b‐strands are oriented perpendicularly to the fibril axis (Sunde and Blake, 1997). Electron microscopy (EM) and atomic

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force microscopy (AFM) revealed that amyloid fibrils possess diverse morphologies at the fibrillar level. Single protofilaments can be straight or curved, with a diameter of 2–5 nm, showing no helical twist. Fibrils usually consist of 2–6 protofilaments, twisting together in a rope‐like or ribbon form with a diameter of 7–15 nm (Serpell et al., 2000). High‐resolution structure determination of amyloid fibrils is a grand challenge. Traditional spectroscopic methods fail because of the insoluble nature or the large size of the fibrils. Crystallization for X‐ray is complicated because fibrils favor growth in one direction. The crystal structure of the amyloid form of the GNNQQNY peptide has recently been solved (Nelson and Eisenberg, 2006). An extended b‐sheet is formed in the crystal, with each of the b‐strands consisting of a single peptide. The structure revealed a tight packing of the side chains between two b‐sheets, excluding water molecules. Solid‐state NMR spectroscopy may provide high‐resolution information on amyloid structure. Tycko (2006) and coworkers built a model structure of the Ab(1–40) amyloid b peptide, associated with Alzheimer’s disease, based on solid‐state NMR constraints. In the model, the different Ab molecules are stacked on each other in a parallel arrangement and in register, forming two b‐sheets. Every single Ab molecule contributes to two b‐strands, one in each b‐sheet. Without providing detailed structural information, hydrogen–deuterium exchange methods combined with NMR spectroscopy, as well as limited proteolysis with mass spectrometry, are capable of collecting site‐specific information on the extent of the rigid amyloid core (Hoshino et al., 2002; Myers et al., 2006a). The morphology of amyloid fibrils grown in vitro highly depends on solution conditions such as buffer composition, pH, temperature, and protein concentration. Even under the same conditions, fibrils with different morphologies can be formed from the same polypeptide. Structural studies revealed that this polymorphism of amyloid fibrils is a reflection of different underlying structures at the molecular level (Petkova et al., 2005). AFM and EM have shown a wide variety of protein aggregates depending on the conditions. Proteins may form disordered aggregates, oligomers, spherical aggregates, prefibrillar aggregates, and fibrils with various morphologies (Kad et al., 2003; Stine et al., 2003). > Figure 10-10 shows a unified view of the various types of structures that can be formed by polypeptide chains in vivo or in vitro.

11.3 Mechanism of Amyloid Fibril Formation The kinetics of amyloid formation and the amyloid content of a protein solution can be studied by light scattering, thioflavin T fluorescence, or other spectroscopic methods. Amyloid formation is usually a nucleation‐dependent reaction showing an initial lag phase, followed by rapid growth. Clearing the solution of any aggregated material by ultracentrifugation prior to the reaction may significantly increase the lag phase. Oppositely, addition of a small amount of preformed aggregated material can reduce or eliminate the lag phase. These observations suggest that the lag phase is the time required for proper nucleation. Under some conditions, no lag time is observable, suggesting that nucleation is not always the rate‐limiting step (Uversky et al., 2002; Pedersen et al., 2004). To understand the in vivo mechanism of amyloidoses, it is important to understand the nucleation process and carefully examine the reaction during the lag phase. Oligomers, spherical or chain‐like aggregates, sometimes termed ‘‘protofibrils,’’ forming prior to fibril formation have been observed in many systems (Chiti and Dobson, 2006). It is extremely important to understand the mechanism of the formation of these prefibrillar species. Cell culture and in vivo studies have revealed their toxicity for living cells, and they may be involved in the pathogenesis of diseases such as Alzheimer’s (Lue et al., 1999; McLean et al., 1999).

11.4 Amyloid Formation of Globular Proteins One group of amyloid‐forming systems consists of peptides or protein fragments that are natively unfolded in the monomeric form (Alzheimer’s Ab, Parkinson’s a‐synuclein). It is generally accepted that stable globular proteins need to partially unfold to an amyloidogenic intermediate for fibril formation. This is

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. Figure 10-10 A unified view of the major types of structure that can be formed by polypeptide chains

supported by experimental data showing increased amyloidogenicity under conditions that destabilize the native state. Destabilizing mutations may also promote amyloid formation, which explains the mechanism of some hereditary diseases (Raffen et al., 1999; Canet et al., 2002). Moreover, under carefully chosen denaturing conditions, most proteins are capable of aggregation and amyloid formation in vitro, suggesting that the amyloid state is a general property of polypeptide chains (Stefani and Dobson, 2003). However, out of thousands of different proteins in the human body, only some two dozen are responsible for diseases, indicating that protein sequences and the cellular machinery have evolved to avoid unwanted protein aggregation.

11.5 Physicochemical and Sequence Determinants of Amyloid Formation Hydrophobicity of the peptide chain has been shown to influence its aggregation propensity (Otzen et al., 2000). Clusters of consecutive hydrophobic residues are avoided by evolution (Schwartz et al., 2001). Another crucial factor is the charge of the polypeptide chain. A high net charge may prevent aggregation of the polypeptide (Chiti et al., 2002). Secondary structure propensity may also affect amyloid formation: a high propensity to form b‐sheet structure and low a‐helix propensity are likely to increase the probability of aggregation (Chiti and Dobson, 2006). Alternating patterns of polar and nonpolar residues, which promote b‐sheet formation, are less frequent in natural protein sequences than expected on a random basis (Broome and Hecht, 2000). Proline residues break b‐sheet structures, and hence inhibit aggregation (Steward et al., 2002; Parrini et al., 2005). The presence of b‐bulges on b‐strands exposed on the protein surface has been found to protect against the formation of intermolecular b‐sheets (Richardson and Richardson, 2002).

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11.6 Factors Inducing or Inhibiting Amyloid Formation Under Physiological Conditions Amyloid formation may be facilitated under proper in vitro conditions, but in vivo, it always occurs under physiological conditions. Some disease‐related proteins such as b2‐microglobulin cannot form amyloid fibrils at physiological pH in vitro, suggesting the presence of unknown factors contributing to the aggregation process in vivo. Collagen, apolipoprotein‐E, heparin, serum amyloid P component, and low concentrations of sodium dodecyl sulfate have been reported to promote the aggregation of b2‐microglobulin under physiological conditions (Yamamoto et al., 2004; Myers et al., 2006b). Other molecules binding to and stabilizing the native state may prevent oligomerization or amyloid formation of disease‐ related proteins in the human body, and might serve as effective therapeutic agents in the future.

12

New Biophysical Techniques for the Study of Protein Folding

In the last decade a wide range of physical and chemical methods complemented the fundamental techniques to study protein folding. We give a brief summary of the advances of these methods.

12.1 Rapid Mixing Methods The mechanism of protein folding can be studied by two different groups of approaches. Equilibrium methods provide information about possible folding intermediate states or deduce rate constants from the molecular fluctuations or dynamic properties of the system. Relaxation methods follow the change of the system evolving toward a new equilibrium after a rapid perturbation of its extrinsic variables, such as temperature, pH, pressure, or solvent composition (Roder et al., 2004). The time required for folding varies greatly among proteins, ranging from microseconds to minutes (Roder and Shastry, 1999; Roder et al., 2006). The smallest protein molecules with no folding intermediates fold on the microsecond timescale, which, on one hand, might make them suitable for in silico‐folding simulation studies. On the other hand, such fast reactions make the experimental detection of events during the folding process difficult. Basic techniques in the study of folding kinetics are stopped‐flow fluorescence spectroscopy and stopped‐flow circular dichroism. These techniques are capable of monitoring the formation of secondary and tertiary structures during the folding reaction with millisecond time resolution. In such experiments, rapid processes occurring within the dead time of the measurement were observed for many proteins. The challenge to resolve this initial burst phase and to reveal the structural changes taking place during the first millisecond stimulated the development of new, rapid kinetic techniques capable of triggering and monitoring the folding process on the sub‐ millisecond timescale. While the conventional stopped‐flow apparatus, in which a small volume of a freshly made mixture containing the reacting components is injected into the measurement cell, is quite economical and offers a wide range of applications (Gibson and Milnes, 1964), its dead time is usually about 1 millisecond or longer. Continuous‐flow methods extend the time resolution to the microsecond time range (Chan et al., 1997; Takahashi et al., 1997; Shastry et al., 1998; Akiyama et al., 2002). In the continuous‐flow cell, solutions are mixed under highly turbulent conditions to achieve complete mixing. The kinetics of the reaction is monitored under steady‐state flow conditions as a function of the distance downstream from the mixer by using relatively simple and inexpensive detection methods. Using this technique, it has become possible to study the initial collapse and formation of intermediates in the early stage of the folding reaction, during the burst phase (Uzawa et al., 2004; Welker et al., 2004; Kimura et al., 2005).

12.2 Real‐Time NMR Spectroscopy Nuclear magnetic resonance (NMR) spectroscopy has greatly contributed to our understanding of the protein‐folding problem. NMR provides high spatial resolution and a broad timescale ranging from

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picoseconds to days. Hydrogen–deuterium exchange experiments, revealing dynamical events at an atomic level, have illuminated the process of unfolding from the native state and the structure of folding intermediates (Wagner and Wuthrich, 1982; Wand et al., 1986; Bai et al., 1995). The quenched‐flow pulse‐ labeling technique has enabled researchers to study the early stages of protein folding using a conventional NMR instrument (Roder et al., 1988; Udgaonkar and Baldwin, 1988; Radford et al., 1992). NMR studies have characterized the properties of denaturant‐induced equilibrium folding intermediate states such as the molten globule (Arai and Kuwajima, 2000). In equilibrium systems, the rates of conversion between distinct conformational states can be calculated from a line shape analysis of the NMR resonances (Huang and Oas, 1995), and therefore can provide kinetic data on folding. Slow folding reactions such as cis–trans prolyl isomerization can be directly followed by sequential recording of one‐dimensional (1D) NMR spectra (Balbach et al., 1999). This method is particularly useful for discovering intermediates formed at the late stages of the folding process. Using a stopped‐flow device for injection of the protein solution into the NMR tube that already contains the denaturant or the refolding buffer pushes the dead time of mixing below 1 s (Zeeb and Balbach, 2004). One of the first proteins studied by real‐time NMR was a‐lactalbumin (Balbach et al., 1995). 1D‐NOE (nuclear Overhauser effect) experiments revealed the native‐like compactness of the transient molten globule state of a‐lactalbumin (Forge et al., 1999). These experiments also demonstrated that the transient intermediate closely resembles the well‐characterized stable molten globule state formed at low pH. While 1D‐NMR spectra have limited resolution, multidimensional NMR can provide high spatial resolution information on the folding process. Because recording multidimensional spectra is time consuming, only slow processes could be followed directly by sequential recording (Liu et al., 1996). Balbach and coworkers (1996, 1999) developed new methods to reconstruct the kinetic history of folding reactions from a single two‐dimensional NMR spectrum recorded during the entire time course of the reaction. The basis of these methods is that the line widths and intensities reflect the history of the folding events occurring during spectral accumulation. When applied to a‐lactalbumin, the technique demonstrated the cooperative nature of the folding of the main chain.

12.3 Chemically Induced Nuclear Polarization Chemically induced nuclear polarization (CIDNP) can be used to probe the solvent accessibility of certain aromatic residues in proteins (Mok et al., 2003; Mok and Hore, 2004). The reactive collision of polarizable amino acids such as tryptophan, tyrosine, and histidine with a photoexcited dye such as flavin mononucleotide (FMN) results in an electron transfer (in the case of Trp and Tyr) or proton transfer (His) reaction forming a pair of radicals. Electron‐nuclear hyperfine interactions between the two radicals result in a significant enhancement of NMR signals. The ‘‘photosensitizer’’ flavin molecule can be excited by laser as light source. For the photoreaction to take place, the aromatic side chains must be accessible to the photosensitizer, e.g., located on the surface of the protein molecule. The CIDNP spectrum is recorded immediately after the laser flash and corrected by a ‘‘dark’’ spectrum recorded without irradiation. Besides the equilibrium studies of protein surfaces, the technique can be combined with a stopped‐flow apparatus and in this way it can be used to study folding intermediates. Using CIDNP pulse‐labeling technique, the exposed tryptophan and tyrosine residues in a molten globule state can be identified (Lyon et al., 2002; Mok et al., 2003).

12.4 High‐Pressure NMR Spectroscopy When high pressure is applied to a protein solution, it shifts the conformational equilibrium of the protein molecules toward lower volume conformers, thereby decreasing the partial molar volume of the protein. The combination of high pressure with heteronuclear two‐dimensional NMR spectroscopy provides atomic resolution information on the structure of the protein molecule at different stages of the folding process (Kamatari et al., 2004). By varying the pressure, one can explore the conformational space from the folded to the unfolded conformer. In recent years, numerous studies using high‐pressure NMR spectroscopy have

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been carried out (Akasaka and Yamada, 2001) on locally disordered (Kuwata et al., 2001; Kuwata et al., 2002; Kitahara et al., 2005), molten globule (Kitahara et al., 2002; Lassalle et al., 2003), unfolded (Kamatari et al., 2001; Arnold et al., 2002; Refaee et al., 2003) as well as oligomeric or aggregated states of proteins (Niraula et al., 2004; Silva et al., 2006).

12.5 Protein Folding and Dynamics Studied by Mass Spectrometry Mass spectrometry of protein molecules has become a rapidly developing field in the last decade (Konermann and Simmons, 2003; Eyles and Kaltashov, 2004). In comparison with NMR spectroscopy, which provides site‐specific information averaged in time, mass spectrometry is capable of detecting different conformers coexisting in the protein solution. This method is especially useful for the study of low‐ populated intermediate states and is free of the molecular size limitation of NMR spectroscopy. Because of its high sensitivity, a protein concentration in the femtomolar range is sufficient for analysis. Structural and dynamic properties of various conformational states can be studied by hydrogen/deuterium exchange (HDX) combined with mass spectrometry. Recently, Kaltashov and coworkers investigated the conformational ensemble of the molten globule state of ubiquitin (Hoerner et al., 2005). Using protein ion fragmentation in the gas phase, they evaluated the stability of various segments of the protein in the molten globular state. By the method of pulse‐labeling HDX‐MS, it is possible to study the kinetics of folding and to explore complex folding scenarios with parallel pathways (Konermann and Simmons, 2003). Co‐populated protein conformers can be detected and characterized directly by electrospray ionization mass spectrometry (ESIMS) (Mohimen et al., 2003; Borysik et al., 2004). Protein surface areas in solution may be determined by ESIMS (Kaltashov and Mohimen, 2005). Limited proteolysis with ESIMS provides site‐specific structural information on different conformational states of the protein molecules including protein aggregates and the amyloid state (Myers et al., 2006a).

12.6 Mechanical Unfolding of Proteins In the first studies of the mechanical unfolding of single protein molecules using AFM, the giant sarcomeric protein titin, consisting of a large number of immunoglobulin segments, was used (Erickson, 1997; Rief et al., 1997; Rounsevell et al., 2004). Because of the heterogeneity of titin domains, it was not possible to assign the individual force peaks to specific domains. Using tandem repeats of a single domain, constructed by protein engineering techniques, it was possible to explain the mechanical characteristics of single domains in terms of their specific structures (Carrion‐Vazquez et al., 1999; Carrion‐Vazquez et al., 2000). Using force‐measuring optical tweezers, it is possible to induce mechanical unfolding and refolding of individual molecules (Kellermayer et al., 1997). In a recent work, Cecconi and coworkers (2005) showed that E. coli ribonuclease H molecule unfolds in a two‐state manner and refolds through a transient molten globule‐like intermediate. We may expect significant progress in the application of other new techniques such as the study of single‐molecule folding kinetics by optical techniques in the near future (Lipman et al., 2003).

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Conclusions

In spite of the great advances in experimental technique and the tremendous boost in computational power we have witnessed in the past decades, the protein‐folding problem is still far from solved. We already have a good understanding of some of the simpler protein‐folding mechanisms, and we can simulate the folding of a few, very small proteins. However, we are still at a loss when the folding behavior of more complex, multidomain proteins is to be explained, especially when protein–protein interactions, misfolding, aggregation, and amyloid formation complicate the situation—such as in a living cell. Although we have known

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since Anfinsen that sequence determines structure (in a given environment), and the field of protein structure prediction has made great progress, we are still not, in general, capable of predicting the structure from a sequence. The research of protein folding, however, is no longer an exotic field with only theoretical significance. Diseases such as Alzheimer’s or Parkinson’s have taught us that protein folding can go awry, and misfolded or aggregated proteins can cause a lot of trouble. In order to successfully defend us against these and other ‘‘folding diseases,’’ we should reach a level of understanding of folding phenomena that is not just descriptive but allows us to influence the folding behavior of proteins in the way we want. Therefore, the research of folding continues, and in all likelihood will bring about great advances in the development of new experimental techniques and new theoretical approaches.

Acknowledgments A.S. and J.K. were supported by Bolyai Jańos fellowships. A.S. and P.Z. were supported by grants from the Hungarian Scientific Research Fund (OTKA T046423 and NI‐61915). S.O. was supported by a grant from the Hungarian Scientific Research Fund (OTKA D‐38480).

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T. Kudo . M. Takeda

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346

2 2.1 2.2 2.3 2.4 2.5

Inclusion Bodies in Neurodegenerative Dementias . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346 Amyloidopathy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346 Tauopathy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 347 Synucleinopathy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349 Polyglutamine Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349 Prion Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 350

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Mechanism of UPS Proteolysis and Neurodegenerative Dementia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 350

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Relationship Between UPS and Endoplasmic Reticulum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 352

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Neurodegenerative dementias involving aberrant protein aggregation

Abstract: Inclusion bodies are common features of neurodegenerative disorders. Recently, it was shown that inclusion bodies are the result of aberrant protein aggregation. Thus, neurodegenerative diseases are classified into five categories: amyloidopathy, tauopathy, synucleinopathy, polyglutamine diseases, and prion diseases. Because the aggregated proteins are insoluble and highly ubiquitinated, the proteins that should have been metabolized by the ubiquitin‐proteasome system (UPS) accumulate to form inclusion bodies. We propose the following mechanisms as a hypothesis for neurodegenerative dementia: (1) aberrant protein aggregation overloads the UPS, (2) the disturbed UPS affects endoplasmic reticulum‐associated degradation (ERAD), (3) disturbed ERAD impairs the unfolded protein response (UPR), and (4) prolonged UPR causes ER‐mediated apoptosis. List of Abbreviations: AD, Alzheimer’s disease; APH-1, anterior pharynx defective-1; APP, amyloid precursor protein; ASK 1, apoptosis-signaling kinase 1; ATF6, activating transcription factor 6; Aβ, β-amyloid; BACE 1, β-site APP cleaving enzyme 1; BCL2, B-cell CLL/lymphoma 2; BiP, immunoglobulin heavy-chain binding protein; BSE, bovine spongiform encephalopathy; CHOP, CCAAT/enhancer binding protein; CJD, Creutzfeldt-Jacob disease; DRPLA, dentatorubral-pallidoluysian atrophy; E1, ubiquitinactivating enzyme; E2, ubiquitination enzyme; E3, ubiquitin ligate; eIF2α, eukaryotic translation initiation factor 2α; ER, endoplasmic reticulum; ERAD, ER-associated degradation; FTDP-17, front temporal dementia and parkinsonism linked to chromosome 17; GSK-3, glycogen synthase kinase-3; IRE 1, inositolrequiring kinase 1; IVS, intervening sequence; JNK, c-jun amino terminal kinase; LTP, long-term potentiation; MJD, Machado-Joseph disease; PEN-2, presenilin enhancer-2; PERK, PKR-like ER-associated kinase; PKR, protein kinase regulated by RNA; PrP, prion protein; PS 1, presenilin 1; S1P, site-1 protease; S2P, site-2 protease; SBMA, spinal and bulbar muscular atrophy; SCA, spinocerebellar; TRAF2, tumor necrosis factor receptor-associated factor 2; UPR, unfolded protein response; UPS, ubiquitin-proteasome system; XBP-1, X-box-binding protein-1

1

Introduction

Dementia occurs due to a variety of causes, leading to the classification of either dementias occurring from the central nervous system itself such as neurodegenerative dementias, or secondary dementias occurring from other events not affecting the central nervous system. The causes of most neurodegenerative dementias are still unknown. However, they all share a common feature, i.e., gradual neuronal death in various brain regions resulting in a neurodegenerative disorder. Specific inclusion bodies are observed in lesioned brains of most neurodegenerative disorders suggesting that they are the products of a common pathological process. Recently, it was shown that inclusion bodies are the result of aberrant protein aggregation. Studies on the mechanisms involved in aberrant protein aggregation have become the main topic of interest in research related to neurodegenerative disorders.

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Inclusion Bodies in Neurodegenerative Dementias

2.1 Amyloidopathy Inclusion bodies are common features of neurodegenerative disorders. Alzheimer’s disease (AD), a representative degenerative dementia, is characterized neuropathologically by senile plaques and neurofibrillary tangles consisting of b‐amyloid (Ab) proteins and highly phosphorylated tau proteins, respectively. Recently, the amyloid cascade hypothesis has been overwhelmingly approved as the cause of AD pathology (Hardy and Selkoe, 2002). It has been suggested that the process of AD pathology consists of (1) increase of Ab in the brains, (2) amyloid depositions, (3) neurofibrillary changes, and (4) neuronal death. Namely, cerebral Ab accumulation is thought to first affect the brains of patients with AD resulting in neurofibrillary changes as a subsequent event through several steps (> Figure 11-1). Therefore, AD is classified as an amyloidopathy.


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. Figure 11-1 Amyloid cascade hypothesis (Hardy and Selkoe, 2002). The pathological sequence of Alzheimer’s disease is explained by the amyloid cascade hypothesis. It shows that an increase in the levels of Ab42 is the critical step in the pathology

Ab is generated from amyloid precursor protein (APP). Recent studies revealed that APP is first cleaved by the b‐secretase BACE 1, at the N terminus of Ab, followed by an intramembranous second cleavage by g‐secretase at the C terminus of Ab. g‐Secretase cleaves APP indiscriminately, resulting in Ab40 and Ab42 among other Ab peptides (> Figure 11-2). Ab42 is more prone to aggregation compared to Ab40. The activity of g‐secretase is controlled by the PS1 complex, which consists of heteromeric molecules including nicastrin, PEN‐2, and APH‐1 (Haass, 2004). Although PS1 is mainly located in the endoplasmic reticulum (ER), g‐complexes have also been demonstrated in the plasma membrane and endosomes where they execute g cleavage (Annaert et al., 1999; Cupers et al., 2001). Ab exists in several different physical states, including monomers, oligomers, or fibrils. Among these states, Ab oligomers are a very potent toxic species, as even nanomolar concentrations have been shown to kill mature neurons (Lambert et al., 1998). Moreover, Ab oligomers appear to interfere with many critical neuronal activities, including long‐term potentiation (LTP) (Lambert et al., 1998). There are some objections to the amyloid cascade hypothesis. One of the concerns with the hypothesis is that the number of amyloid deposits in the brain does not correlate well with the degree of a patient’s cognitive impairment. In contrast, Braak et al reported that the pathological stage of AD can be classified by the distribution of neurofibrillary tangles (Braak and Braak, 1991). In this sense, AD can also be classified as a tauopathy.

2.2 Tauopathy Tauopathy refers to disorders characterized by neurofibrillary tangles composed of highly phosphorylated tau in neurons or glia. Frontotemporal dementia, corticobasal degeneration, and progressive supranuclear

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. Figure 11-2 Generation of Ab from APP. bAPP, located through the membrane, is cut at three enzymatic cleavage sites: a cutting, b cutting, and g cutting sites. a cutting and g cutting generate P3, whereas b cutting by BACE and g cutting generate Ab40 or Ab42

palsy, as well as AD, belong to this category and neurofibrillary tangles are commonly observed in neurological disorders involving neuronal loss. Tau is a microtubule‐associated protein that binds and stabilizes microtubules. Six isoforms of human tau are generated by alternative splicing based on the inclusion or absence of exons 2 and 3 in the N terminus and the inclusion or absence of exon 10 in the C terminus. Isoforms containing exon 10 have two types of tubulin‐binding domains: three‐repeat tau and four‐repeat tau (> Figure 11-3). According to the amyloid cascade hypothesis, in AD the phosphorylation of tau has been regarded as just a consequence of amyloidopathy. However, studies of FTDP‐17 (frontotemporal dementia and parkinsonism linked to chromosome 17) showed that abnormalities of tau itself cause neurofibrillary changes and neuronal loss (Foster et al., 1997). FTDP‐17 is characterized by autosomal dominant inheritance. Tau gene mutations specific to this disorder are located in exon 1, exons 9–13, and in the stem–loop structure of the intron following exon 10 (IVS 10). To date, more than 30 different mutations have been reported. Neuropathological studies have revealed a tendency for patients with mutations in IVS 10 or exon 10 (ex. N279K) to exhibit deep gray matter lesions resembling those of progressive supranuclear palsy or corticobasal degeneration, and marked fibrillary changes in both neurons and glia (Reed et al., 1998; Goedert et al., 1999). On the other hand, other tau mutations in patients show frontotemporal degeneration with prominent neurofibrillary changes. However, certain mutations exhibit a considerable variety of neuropathological features among patients with the same mutation. The highly phosphorylated tau in neurofibrillary tangles is thought to result from an imbalance between kinases and phosphatases. There have been several reports on the role of kinases and phosphatases in regard to neurofibrillary tangle formation. Among them, an important role for glycogen synthase kinase‐3 (GSK‐3) in the formation of neurofibrillary tangles is becoming clearer, especially in response to Ab


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. Figure 11-3 Isoforms of human tau. There are six isoforms of human tau produced by alternative splicing in the insertion and tubulin‐binding domains

(Phiel et al., 2003). Inhibition of GSK‐3 prevents neurofibrillary tangle formation by preventing the phosphorylation of tau. However, GSK‐3 activation alone does not explain all of the phosphorylation sites on tau seen in neurofibrillary tangles (Lucas et al., 2001). It is assumed that GSK‐3 induces highly phosphorylated tau in cooperation with other activated kinases and inactivated phosphatases.

2.3 Synucleinopathy In synucleinopathy, Lewy bodies consisting of a‐synuclein are observed in neurons. Diffuse Lewy body disease and Parkinson’s disease are classified in this category. Abnormal inclusions containing a‐synuclein appear in glia and neurons. a‐synuclein is present in the presynaptic terminus and nuclei. The a‐synuclein gene was identified as the causative of familial Parkinson’s disease in South Italy (Polymeropoulos et al., 1997). Primary a‐synucleinopathies include Parkinson’s disease, Lewy body disease, multiple system atrophy, and neurodegeneration with brain iron accumulation type 1. Lewy bodies and/or intracytoplasmic a‐synuclein aggregates also occur in various tauopathies, categorized as secondary a‐synucleinopathies; neuronal a‐synuclein inclusions are frequently found in the amygdala of patients with Alzheimer’s disease, Down’s syndrome with Alzheimer pathology, parkinsonism–dementia complex of Guam, diffuse neurofibrillary tangle with calcification, and neuroaxonal dystrophy with neurofibrillary alteration.

2.4 Polyglutamine Diseases Polyglutamine diseases, such as Huntington’s disease, are disorders with abnormally extended polyglutamine chains due to the expansion of the CAG repeat in a causal gene. The encoded proteins containing abnormally extended polyglutamine chains aggregate in intraneuronal inclusions. To date, nine hereditary neurodegenerative diseases: Huntington’s disease, spinal and bulbar muscular atrophy (SBMA), dentatorubral‐pallidoluysian atrophy (DRPLA), Machado–Joseph disease (MJD), spinocerebellar ataxia type 1 (SCA1), type 2 (SCA2), type 6 (SCA6), type 7 (SCA7), and type 17 (SCA17), have been reported. All of the diseases display autosomal dominant heredity except SBMA, which shows X‐linked heredity. There is no

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homology between causal genes except in their polyglutamine chains. Therefore, it has been speculated that expanded polyglutamine chains cause neuronal damage. Polyglutamine chain‐repeat length increases between generations. If the number of repeats crosses a threshold, the causal genes are unstable and repeat length increases, especially in paternal transmission. As repeat length increases, the age of onset becomes younger and the symptoms worsen. This phenomenon is termed anticipation (Margolis and Ross, 2003).

2.5 Prion Diseases Prion means a proteinaceous infectious particle: it was found in the infectious fraction resistant to detergents and proteinases extracted from the brains of goats with scrapie. Prion protein (PrP) is encoded by a gene endogenous to the host and predominantly expressed in the central nervous system. Infectious PrP (PrPSC) is rich in b‐sheet structures and has a core resistant to proteinases, whereas normal PrP (PrPC) is not infectious and degradable by proteinases. It has been speculated that PrPSC seeds the conversion of PrPC into PrPSC to generate a polymer of PrPSC (Prusiner, 1998). The accumulation of this polymer causes progressive abnormalities of neurons. Prion diseases are classified into idiopathic diseases, contagious diseases, and hereditary diseases. While sporadic Creutzfeldt–Jacob disease (CJD) is idiopathic, subtypes include iatrogenic CJD, such as caused by dura grafting, and variant CJD, which was recently suspected of being linked to bovine spongiform encephalopathy (BSE).

3

Mechanism of UPS Proteolysis and Neurodegenerative Dementia

Generally, inclusion bodies seen in neurodegenerative disorders are composed of unfolded and insoluble proteins that are highly ubiquitinated. Considering this fact in neurodegenerative disorders, the proteins that should have been metabolized by the ubiquitin‐proteasome system (UPS) accumulate, for some unknown reason, to form inclusion bodies. UPS proteolysis begins with the binding of ubiquitin to the e‐amino group of a lysine residue on a target protein with covalent bonds, in an ATP‐dependent manner. The ubiquitin‐activating enzyme (E1) first recruits ubiquitin and passes it to the ubiquitination enzyme (E2). Then, E2 and the target protein bind to ubiquitin ligase (E3) to allow ubiquitin to bind to the target protein. By repeating this reaction, ubiquitin is sequentially added to the target protein to form a polyubiquitin chain. Target proteins containing this polyubiquitin chain (more than four ubiquitin molecules) are recognized by the 26S proteasome and selectively degraded. Prior to degradation in the proteasome, ubiquitins are removed from the target proteins for recycling by a deubiquitin enzyme, and target proteins are cleaved into small peptides (> Figure 11-4). Intervention of the UPS was clarified by studies of familial degenerative dementia due to abnormal genes. Parkin, a causal gene of early onset autosomal recessive inheritance‐related Parkinson’s disease, was identified as an E3 enzyme. In Parkinson’s disease, the metabolism of Pael receptors by the proteasome is blocked leading to accumulation of the receptors in dopaminergic neurons and neuronal death (Kitada et al., 1998; Imai et al., 2001). In some heritable degenerative dementias, gene mutations cause aggregation of proteins that inhibit normal proteasomal function, possibly leading to neuronal death. Pathogenesis of FTDP‐17 is thought to involve decreased metabolism of mutant tau by the proteasome (Foster et al., 1997). A report concerning familial Parkinson’s disease investigated the involvement of a‐synuclein accumulation in Lewy body formation and the pathological process of Parkinson’s disease. Huntington’s disease displays autosomal dominant inheritance, characterized by extension of the CAG repeat in Huntingtin (Nishitoh et al., 2002). It was shown that extended polyglutamine chains can also decrease proteasome function. As mentioned above, aggregation of aberrant proteins leading to inclusion formation, due to dysfunction of the UPS, is a common process in neurodegenerative dementias.


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. Figure 11-4 Ubiquitin proteasome system (UPS). The ubiquitin‐activating enzyme, E1, uses ATP to activate ubiquitin (Ub) to a higher energy. The activated ubiquitin is transferred onto conjugating proteins, E2, and then covalently linked to a protein substrate bound to a specific ubiquitin protein ligase, E3. There are multiple classes of E3s, each one recognizing a specific motif on the substrate. The substrate bound to E3 is polyubiquitinated via the successive addition of activated ubiquitins to internal Lys residues of the previously conjugated ubiquitin to prepare proteins for degradation by the proteasome

4

Relationship Between UPS and Endoplasmic Reticulum

Why does dysfunction of the UPS, due to aggregation of abnormal proteins, cause neuronal death/ apoptosis? The ER is the organelle responsible for controlling protein folding and assembly by expressing molecular chaperones and is the site of other quality control systems for proteins. Thus, various stresses to the ER (ER stress) cause accumulation of unfolded or misfolded proteins in the lumen of the ER. Because this situation disrupts ER homeostasis, the ER has developed highly specific signaling pathways generally termed the ‘‘unfolded protein response’’ (UPR). The UPR consists of three stages. First, when mammalian cells are subjected to ER stress, the immediate response is the activation of PKR‐like ER‐associated kinase (PERK), which inhibits protein biosynthesis through phosphorylation of eukaryotic translation initiation factor (eIF2a) (Harding et al., 1999). Second, transcriptional programs are altered through IRE1 and XBP‐1 leading to induction of ER chaperones, such as BiP, for the refolding of unfolded proteins (Yoshida et al., 2001). Upon ER stress, ATF6 transits to the Golgi compartment where it is cleaved by S1P and S2P proteases to yield a cytosolic fragment (Haze et al., 1999). The free ATF6 fragment migrates to the nucleus to activate transcription. Third, unfolded or misfolded proteins in the ER lumen are retrotranslocated to the cytoplasm, where they are ubiquitinated and degraded by the proteasome. This process is called endoplasmic reticulum‐associated degradation (ERAD) (Werner et al., 1996). Via ERAD, there is a linkage between ER and proteasome function. Thus, it is speculated that proteasome dysfunction may cause UPR followed by ER‐mediated apoptosis as described below (> Figure 11-5). If an overload of unfolded or misfolded proteins in the ER is not resolved, prolonged UPR activation leads to apoptosis. Several previous studies have analyzed apoptosis‐related molecules centered on the mitochondria. However, it has recently been reported that some apoptotic processes originate in ER. The three known proapoptotic pathways emanating from the ER are mediated by IRE1, caspase‐12, and CHOP, respectively (> Figure 11-5). Under ER stress, activated IRE1 recruits the cytosolic adapter TRAF2 to the ER membrane (Yoneda et al., 2001). TRAF2 activates the apoptosis‐signaling kinase 1 (ASK1) to activate c‐Jun

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. Figure 11-5 The unfolded protein response

amino terminal kinase (JNK) for apoptosis (Urano et al., 2000). Caspase‐12 is an ER‐associated proximal effector of the caspase activation cascade and activates caspase‐3, leading to apoptosis (Nakagawa et al., 2000). Because caspase‐12 has been reported to be expressed in mice and rats, we screened a human colon cDNA library using the sequence of mouse caspase‐12 as a probe and detected caspase‐4 with a high frequency. We also showed that caspase‐4 was localized in the ER membrane and was strictly activated by ER stress inducers. Moreover, it was demonstrated that expression of caspase‐4 was increased in brains of patients with AD. These findings suggested that caspase‐4 may function as an ER stress response caspase in humans, and may be involved in neuronal death in AD (Hitomi et al., 2001). A third death‐signaling pathway activated by ER stress is mediated by transcriptional activation of genes encoding proteins with proapoptotic functions. Activation of the UPR transducer PERK results in activation of the transcription of CHOP, a b‐ZIP transcription factor that potentiates apoptosis, possibly through repressing the expression of the apoptotic repressor BCL2 (Ma et al., 2002).

5

Conclusion

In most neurodegenerative dementias, pathogenesis remains unclear, but characteristic inclusion bodies consisting of aberrant protein aggregation are observed. Recently, it has been clarified that these aberrant aggregations commonly involve the UPS. It was also shown that the UPS is related to the UPR through ERAD. Therefore, in this chapter, we propose a hypothesis for the mechanism of neurodegenerative dementia as follows: (1) aberrant protein aggregation overloads the UPS, (2) the disturbed UPS affects ERAD, (3) disturbed ERAD impairs the UPR, and (4) prolonged UPR causes ER‐mediated apoptosis. On the basis of this hypothesis, we can devise therapeutic strategies for each step of the neurodegeneration.


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Polyglutamine Diseases

H. Okazawa

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 356

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Clinical Features of Polyglutamine Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 356

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7

Causative Gene and Functions of the Product . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357 Huntingtin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357 Androgen Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357 Ataxin‐1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357 Ataxin‐2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358 Ataxin‐7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358 TATA‐Binding Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358 DRPLA Product (Atrophin‐1) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358

4 4.1 4.2 4.3 4.4 4.5 4.6

Metabolism of Abnormal Polyglutamine Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 Processing of Polyglutamine Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 Basic Mechanisms of Aggregation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 360 Ubiquitination of Polyglutamine Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 360 Proteasome‐Mediated Degradation of Polyglutamine Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 361 Nuclear Transport of Polyglutamine Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 361 Aggregation Sites in the Cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 361

5 5.1 5.2 5.3 5.4 5.5

Cellular Dysfunction by Polyglutamine Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362 Transcriptional Disturbance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362 ER Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362 Proteasome Dysfunction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Mitochondrial Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Which Causes Symptoms, Neuronal Death or Neuronal Dysfunction? . . . . . . . . . . . . . . . . . . . . . . . . . . . 363

6 6.1 6.2 6.3 6.4 6.5 6.6 6.7

Frontier of Therapeutic Approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Aggregation Modifiers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 Cross‐Linking Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 Cleavage Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 Transcriptional Upregulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 RNAi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 Modification of Nuclear Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366 Drugs for Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366

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Polyglutamine diseases

Abstract: Polyglutamine diseases are a major group of neurodegeneration induced by mutant proteins containing an expanded polyglutamine tract sequence. In this chapter, recent findings on aggregation process and metabolism of mutant proteins are summarized. In addition, the development of molecular therapeutics against polyglutamine diseases are discussed.

1

Introduction

The concept of ‘‘polyglutamine disease’’ originates from the discovery of the causative gene of Kennedy’s disease (spinobulbar muscular atrophy) by the group of Dr. Fischbeck (Laspada e al., 1991). They described that a new type of mutation in the androgen receptor gene, which is not point mutation, deletion, or translocation but expansion of the CAG triplet repeat, is linked to the onset of this motor neuron disease. In this case, mutant androgen receptor possesses a longer CAG repeat sequence expanded more than 40 repeats, which results in a mutant protein containing an abnormally long polyglutamine tract. Transgenic mouse models of Huntington’s disease and spinocerebellar ataxia 1 expressing the expanded polyglutamine tract confirmed that the poly‐Q tract expansion is the primary cause of neurodegeneration. It was surprising that CAG repeat sequences, distributed widely in the genome and used for genome markers for genetics study, are the direct cause of diseases. The new concept of mutation stimulated subsequent discoveries of new disease genes. For example, elongation of another type of triplet repeat (GCG/Ala repeat) was found in poly(A)‐binding protein nuclear 1 (PABPN1) as the cause of oculopharyngeal muscular atrophy (Brais et al., 1998). In addition, elongation and shortening of dinucleotide repeat in poly‐Q‐ binding protein‐1 (PQBP‐1) (Karlscheuer et al., 2003) have been shown to cause mental retardation. At the beginning of the triplet repeat disease research, it was discussed whether CAG repeat expansion causes neuronal cell death through production of an abnormal protein or through certain genomic effects such as transcriptional silencing of neighboring genes. However, in the case of poly‐Q repeat diseases (exon‐coded CAG repeat disorders), it is now believed that abnormal proteins directly induce the disease phenotype. Furthermore, studies on polyglutamine diseases have unraveled new pathophysiology such as protein aggregation, endoplasmic reticulum (ER) stress, mitochondrial stress, nuclear transport, and transcriptional repression. Although their relationship was not yet clear, recent data suggest that these critical pathological processes are linked mutually. Collectively, polyglutamine diseases are a new, but most well–investigated, neurodegeneration associated with protein aggregation process. In this chapter, I describe polyglutamine diseases by focusing on the metabolism and function of pathogenic proteins.

2

Clinical Features of Polyglutamine Diseases

So far, nine neurodegenerative disorders (Huntington’s disease, Kennedy’s disease, SCA1, SCA2, SCA3, SCA6, SCA7, SCA17, and DRPLA) are known to belong to polyglutamine diseases in which abnormal expansion of poly‐Q repeats is linked to the disease onset (see reviews by Gusella and MacDonald, 2000; Zoghbi and Orr, 2000; Ross et al., 2002; Taylor et al., 2002; Bates and Hockly 2003; Okazawa, 2003). The number of expanded repeats is somewhat variable among different disorders, whereas generally speaking repeats more than 40 lead to abnormal phenotype. The critical number is now known to be the threshold of protein conformational change (Scherzinger et al., 1999), which is essential for producing the most important pathological hallmark, inclusion body. In both human and mouse models, it is known that inclusion body grows during the course of the disorders. It was previously believed that inclusion body in neurons directly leads to cell death. However, the idea is now challenged by a number of recent data. For example, human pathological data suggest that inclusion bodies are formed in the region not remarkably affected and that neurons can survive for a long period carrying inclusion bodies. On the basis of affected regions in the brain, polyglutamine diseases can be classified into three groups. In most polyglutamine diseases, cerebellum is the most vulnerable region in the brain. The first group has been diagnosed as spinocerebellar degeneration in classical clinical neurology before DNA diagnosis became available. At the microscopic level, it has been known that there are minor differences in the pathology of


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the cerebellum. For example, although loss of Purkinje cells is remarkable, granule cells are not usually affected in SCA1. However, granule cells are also affected in SCA3. The second group shows motor neuron disease phenotype. Kennedy’s disease, in which lower motor neurons of the spinal cord are predominantly affected, is included in this group. The patient shows muscular atrophy and weakness of limb, trunk, and facial muscles. Some patients of SCA1 show remarkable muscular atrophy and motor neuronal loss in the spinal cord. Thus, SCA1 could be considered to belong to both group 1 and 2. The third group involves Huntington’s disease and maybe SCA17. These diseases mainly affect the basal ganglia and the cerebral cortex, thus patients show involuntary movements and dementia. Dentatorubral‐pallidoluysian atrophy (DRPLA) also belongs to this category.

3

Causative Gene and Functions of the Product

Functions of representative polyglutamine disease gene products are reviewed in this section. Disease proteins share polyglutamine tract sequence, whereas their physiological functions are variable.

3.1 Huntingtin So far, physiological functions of huntingtin (htt) have not been completely understood. However, recent notions from htt‐interacting molecules such as huntingtin‐associated protein (HAP1) and htt‐interacting protein 1 (HIP1) help us to speculate its functions. HAP, which was cloned by two‐hybrid screening, is predominantly expressed in the brain and in neurons. It is known to associate with p150Glued, a subunit of dynactin complex (Engelender et al., 1997; Gutekunst et al., 1998). Dynactin complex associates with dynein, an anterograde kinetomotor protein moving on microtubules, and transport vesicles. Recently, it was shown that BDNF‐containing vesicles are carried by the complex, including HAP1 (Gauthier et al., 2004). Furthermore, htt protein also associates with the complex. In the case of mutation, the association between the transport complex and microtubules is reduced, and therefore the transport of BDNF is decreased. The reduction of transport and resultant release of BDGF might be relevant to the pathology. The second molecule HIP, htt‐interacting protein 1, was also cloned by two‐hybrid screening (Kalchman et al., 1997; Wanker et al., 1997). HIP is known to interact with clathrin and a‐adaptin‐binding protein, which are submembranous coating proteins of endocytotic vesicles (Waelter et al., 2001). This finding suggests that htt is located also around endocytotic vesicles. Although functions implicated by HAP and HIP are not identical, it is interesting that they share a similarity, i.e., their function in vesicular transport. Whole scheme of htt functions will be understood more clearly in the future. Htt gene knockout leads to malformation of the nervous system (White et al., 1997) suggesting that Htt protein is involved in neural development or in essential neuronal functions. The finding is consistent with the speculated molecular functions of htt protein.

3.2 Androgen Receptor Androgen receptor is one of the nuclear steroid hormone receptors carrying the poly‐Q tract in its N‐terminal region. Although the expansion was reported to repress transcription directly (Mhatre et al., 1993), the role in aggregation is generally considered to be much more important. However, this question actually remains open because poly‐Q tract in certain transcription factors acts as a transactivation domain (Tanaka et al., 1994). Since physiological functions of the androgen receptor are numerous, I will not go further in this rather short chapter focusing on poly‐Q diseases.

3.3 Ataxin‐1 Ataxin‐1 is clearly a nuclear protein that, even in a normal form, composes the nuclear body. The nuclear body is known to include PML (Skinner et al., 1997), a scaffold of multiple nuclear proteins involved in

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transcription, RNA modification, and cell signaling. A part of ataxin‐1 nuclear bodies colocalizes with other marker proteins of nuclear bodies, such as PQBP‐1 (Okazawa et al., 2002) and LANP (Matilla et al., 1997), whereas the three proteins, PML, LANP, and PQBP‐1 do not completely overlap in all nuclear bodies. It might suggest that ataxin‐1 dynamically moves among different functional fields. LANP is known to be a component of INHAT that is involved in transcription and histone acetylation (Seo et al., 2001). Yue et al. (2001) reported that ataxin‐1 has RNA‐binding activity. PQBP‐1 is a connector protein of RNA polymerase II large subunit C terminus and RNA splicing protein U5‐15kD (Okazawa et al., 2002). These findings suggest that ataxin‐1 is relevant to basic nuclear functions including transcription and also RNA modification.

3.4 Ataxin‐2 In SCA2 caused by mutation of ataxin‐2 gene, although some brain stem neurons possess nuclear inclusions, nuclear inclusions are rare in cerebellar neurons that are dominantly affected in SCA2. It is possible to speculate either that nonaggregated ataxin‐2 in the nucleus is more toxic than aggregated form or that cytoplasmic inclusion found in affected regions is more toxic than nuclear inclusion (Hunyh et al., 2000). In the case of ataxin‐2, its pathological role in the cytoplasm is considered to be more critical than that in the nucleus. Hunyh et al. reported that normal ataxin‐2 protein is located in the Golgi apparatus while mutant ataxin‐2 is not. In the mutant, ataxin‐2 overexpression disrupts the Golgi complex. These results suggest that ataxin‐2 is a critical component of or a relevant molecule to the Golgi complex (Huynh et al., 2003). Meanwhile, it is reported that ataxin‐2 has RNA‐binding motifs to RNA (Shibata et al., 2000). There remains a gap between suggested localization and molecular characteristics suggesting that the function of this molecule may not yet be completely understood.

3.5 Ataxin‐7 SCA7 is clearly different from other types of spinocerebellar degeneration in that patients show retinal degeneration. Although macular degeneration is easy to diagnose, electroretinogram examination reveals widely distributed cone–rod dystrophy of the retina. Correspondingly with the clinical feature, the causative gene product ataxin‐7 binds to CRX, a transcription factor that regulates retinal genes (Laspada et al., 2001; Chen et al., 2004). In addition to the interaction with the specific transcription factor, the yeast ortholog of ataxin‐7, SGF73, turned out to be a component of the SAGA (Spt/Ada/Gcn5 acetylase) complex. Furthermore, a French group revealed that ataxin‐7 is a conserved component of mammalian TFTC/STAGA complex that is required for transcription of a subset of RNA polymerase II‐dependent genes (Helmlinger et al., 2004).

3.6 TATA‐Binding Protein SCA17 is a peculiar disease showing ataxia, short stature, atypical absence, pyramidal signs, and mental deterioration (or dementia). The patients show brain atrophy in the cerebrum, cerebellum, and brain stem. The Tsuji group reported the first case (Koide et al., 1999) and family cases were subsequently reported by the Kanazawa group (Nakamura et al., 2001). The causative gene was known to be the gene for TATA‐ binding protein; the protein recognizes a specific upstream sequence (TATA box) in the promoter region of a gene and recruits RNA polymerase II and other transcription factors.

3.7 DRPLA Product (Atrophin‐1) Expansion of poly‐Q tract sequence in atrophin‐1 (or DRPLA product) leads to degeneration of basal ganglia in the cerebellar and pallidal outflow pathways. Although the function of this protein is not


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completely understood, various data suggest its functions in signaling and transcription. The Yamada group screened binding proteins to the DRPLA product and found IRS p53 and DVL1 as candidate binding partners (Okamura‐Oho et al., 1999), which could activate JNK signaling. They also found that the DRPLA product is phosphorylated by JNK (Okamura‐Oho et al., 2003). Collectively, these results suggest that atrophin‐1 is involved in JNK signaling. On the other hand, a homolog of atrophin‐1 in Drosophila was isolated. It was called atro (or grunge), which acts as a transcriptional corepressor in the earliest stage of embryogenesis (Erkner et al., 2002; Zhang et al., 2002).

4

Metabolism of Abnormal Polyglutamine Proteins

Disease proteins carrying an abnormally expanded polyglutamine tract are translated in the ER. The abnormal proteins tend to have abnormal conformation but chaperone proteins try to unfold and refold their structure. If the trial is unsuccessful, chaperone proteins induce ER stress response. A part of disease proteins in abnormal structures translocate (or leak) to the cytoplasm. They are processed by proteinases and some of them move also to the nucleus. Ubiquitination occurs in these subcellular compartments and ubiquitinated proteins are recognized as the target to be cleared in the proteasome. However, they are accumulated sometimes at the centrosome where factors for protein degradation are concentrated (aggresome). In this case, it is known that aggregated polyglutamine proteins disturb the proteasomal function. A part of the proteins after cleavage moves to the nucleus and forms relatively firm nuclear inclusions. It is not yet clear at which step the polyglutamine peptide is most toxic for neurons. However, recent notions have suggested strongly that monomers or oligomers in abnormal conformations are toxic and not the final aggregates. I will summarize in this section recent findings on the metabolism of polyglutamine proteins (> Figure 12‐1).

. Figure 12‐1 Metabolism of mutant polyglutamine proteins

4.1 Processing of Polyglutamine Proteins It is known that htt, ataxin‐3, ataxin‐6, ataxin‐7, atrophin/DRPLA product, and androgen receptor are cleaved into a peptide containing the polyglutamine sequence (see reviews by Gusella and MacDonald, 2000;

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Zoghbi and Orr, 2000; Ross et al., 2002; Taylor et al., 2002; Bates and Hockly 2003; Okazawa, 2003). It is also highly likely that other polyglutamine disease gene products are also processed. The proteolytic enzymes reported up to now include caspase‐3 for androgen receptor, caspase‐1 for ataxin‐3, calpain family (calpain‐1, ‐5, ‐7, ‐10), and caspase‐3 for htt. The cleaved polyglutamine peptides containing polyglutamine tract were detected in brain samples from patients or in model systems. It is generally believed that shorter polyglutamine peptides are more toxic and more prone to aggregate. Accordingly, some groups reported that inhibition of the cleavage protects animal models from progress of pathology. Meanwhile, it is not always true that cleavage accelerates the pathology. In the case of htt, normal form protein is also coaggregated into inclusion body (Busch et al., 2003). As normal htt plays a protective role for neurons, cleavage of normal protein may be implicated in the pathology. It was not clear whether protein cleavage is similarly performed in the cytoplasm and in the nucleus. However, Lunkes et al. reported recently that nuclear aggregation contained specifically short peptides of about 50kDa cleaved by aspartic endopeptidases while cytoplasmic inclusion body contained longer peptides (Lunkes et al., 2002). The result suggests either that only short peptides could move to the nucleus or that nucleus‐specific endopeptidases cleave htt in the nucleus.

4.2 Basic Mechanisms of Aggregation Peptides containing polyglutamine tract sequence mutually bind and form aggregates in vitro when the repeat length exceeds a disease‐specific threshold. The aggregates formed in cells are called inclusion body, which is a disease‐specific pathological hallmark. Max Perutz proposed that expanded polyglutamines form the polar zipper structure, and it is the most prevailing model in polyglutamine disease research (see review by Perutz, 1996). It is the antiparallel b‐sheet structure stabilized by hydrogen bonds between positive‐ and negative‐charged glutamine side chains. Computer modeling studies suggested that polyglutamine sequences could form other structures such as parallel b‐sheet, b‐hairpin, compact random coil, and compact b‐sheet (Ross et al., 2003). With X‐ray analysis, Perutz and colleagues also suggested that a large b‐helix (20 residues per turn) could be also formed. The Hartl group suggested polymerization of poly‐Q monomers with b-sheet conformation (Sakahira et al., 2002). In the case of htt, in vitro aggregation revealed that the threshold locates between 32 and 37. However, threshold of the repeat number for the onset of human patients is different among polyglutamine diseases. For example, in the case of SCA6, the threshold locates between 18 and 20. Although the reason is not clarified, the difference probably comes from the properties of surrounding sequences.

4.3 Ubiquitination of Polyglutamine Protein Polyglutamine proteins are highly ubiquitinated in neurons. This central dogma is derived from immunohistochemistry of cytoplasmic and nuclear inclusion bodies and from Western blot analysis of intracellular mutant proteins. Meanwhile, it is not so clear which types of cleaved peptides (or noncleaved native proteins) are ubiquitinated. It is also not known whether the abnormal proteins/peptides are ubiquitinated in the nucleus or in the cytoplasm of neurons. Ubiquitin ligase for each polyglutamine protein seems to be different according to the published results. So far, several ubiquitin ligases are known to interact with polyglutamine disease proteins or peptides. The first example is E6‐AP/Ube3a that promotes ubiquitination of ataxin‐1 (Cummings et al., 1999). When Ube3 E3 ubiquitin ligase mutant mice and SCA1 transgenic mice were mated, nuclear inclusion formation was remarkably inhibited and, surprisingly at that time, the pathology of Purkinje cells became much worse. This means that E6‐AP/Ube3 is critically involved in ubiquitination of ataxin‐1. In addition, from the findings, emerged the recently prevailing theory that preaggregates are more toxic for cells. There are some other proteins involved in ubiquitination. Ataxin‐1‐ interacting ubiquitin‐like protein (A1Up) could be involved in the ubiquitination of ataxin‐1 (Davidson et al., 2000) because it contains an ubiquitin‐like region (Ubl), four heat‐shock chaperonin‐binding motifs,


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and an ubiquitin‐associated domain (UBA). The Ubl domain interatcts with S5a subunit of 19S proteasome and the UBA domain binds polyubiquitin chains. These results suggest that A1Up regulates degradation of ataxin‐1 (Riley et al., 2004).

4.4 Proteasome‐Mediated Degradation of Polyglutamine Proteins It has been believed that ubiquitinated polyglutamine proteins or peptides are transported to and degraded by proteasome (Cummings, 1998). However, Venkatraman et al. have shown that small polyglutamine peptides containing expanded repeats are hardly digested by eukaryotic proteasome (Venkatraman et al., 2004). Although their experiments were based on in vitro study, if it is true, we should better assume that ubiquitinated peptides mainly accumulate rather than being cleared from cells. The result might be relevant to the finding that accumulated polyglutamine proteins disturb the function of the proteasome (Waelter et al., 2001). Peptides that could not be digested by the proteasome accumulate around the proteasome complex. The accumulation domain corresponds to the centrosome where breakdown of microtubules are actively conducted. This region is therefore also called the aggresome (Waelter et al., 2001). The result again stimulates the discussion whether ubiquitination protects or damages neurons. If ubiquitination promotes protein degradation or segregation from interacting partners, ubiquitination is supportive for cells, if not, ubiquitination might be potentially toxic for neurons.

4.5 Nuclear Transport of Polyglutamine Proteins In some polyglutamine diseases, the normal form of the disease protein is located in the nucleus. For example, Kennedy’s disease is caused by repeat expansion of a nuclear hormone receptor, androgen receptor. SCA17 is caused by mutation of TATA‐binding protein, an essential nuclear factor binding to TATA box, initiating transcription. Furthermore, mutant forms of most of the polyglutamine disease proteins are definitely transported to the nucleus because they are actually aggregated in the nucleus. However, our knowledge about their nuclear transport remains insufficient in most of the polyglutamine proteins. In the case that normal forms are nuclear proteins the mutant proteins may be transported into the nucleus by ordinary nuclear transport systems through the nuclear pore complex. However, there seems to be some additional factors that accelerate the nuclear import because mutant proteins are accumulated in the nucleus even if the normal forms are located in the cytoplasm. Furthermore, nuclear export of mutant polyglutamine proteins is not yet reported so far.

4.6 Aggregation Sites in the Cell Mutant polyglutamine proteins are aggregated in the cytoplasm and also in the nucleus. The focus of aggregation in the cytoplasm merges on the centrosome (Johnston et al., 1998), which is the organizing center of microtubules for cell division where protein complex formation and protein degradation are actively conducted. Numerous chaperone proteins including Hsp70 and Hsp27 as well as proteasome proteins are located in the centrosome. Mutant proteins are probably transported to centrosome for degradation. However, they would be resistant to degradation by proteasome and overflowed proteins would accumulate in the centrosome. Centrosome embedded by protein aggregation is sometimes called as aggresome. In the nucleus, aggregates are not localized to visible nuclear domains. Instead, they accumulate in nuclear domains that can be detected by specific marker proteins. For example, ataxin‐1 is known to accumulate in PML body and also nuclear body of PQBP‐1. Functional relevance of accumulation in specific nuclear domains is not completely clarified so far as described later.

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Cellular Dysfunction by Polyglutamine Proteins

Mutant polyglutamine disease proteins are known to disturb multiple cellular functions (> Figure 12‐2). ER stress is linked to apoptosis in nonneuronal cells. Proteasomal dysfunction will also trigger the apoptotic cascade if the capacity to refold or degrade abnormal proteins becomes insufficient. It is also reported that mutant proteins associate with mitochondrial membrane. . Figure 12‐2 Relationship between aggregation process of mutant proteins and induced cellular stresses

5.1 Transcriptional Disturbance Transcriptional dysfunction is considered to be a key event in polyglutamine disease pathology. It is because most polyglutamine disease proteins accumulate in the nucleus and are known to interact with many nuclear proteins including transcription factors, transcription cofactors (coactivators and corepressors), and splicing factors (see review Okazawa, 2003; Sugars and Rubinsztein, 2003). Although a few number of transcription‐related factors have been highlighted, it is not known whether transcription of a specific gene mediated by a specific transcription factor is disturbed or general transcription mediated by multiple nuclear factors is abrogated.

5.2 ER Stress Mutant polyglutamine proteins induce ER stress. This response is basically similar to ERAD that protects cells from protein misfolding. It ceases translation, induces chaperone gene upregulation and triggers signaling of apoptosis. The first pathway is mediated by PERK, the second is mediated by IRE1 and ATF6, and the third is mediated by IRE‐JNK cascade or CHOP. These responses basically stop de novo synthesis of toxic proteins and select the pathway that leads to cell suicide when refolding of toxic proteins fail. Therefore, synthesis of unfolded or misfolded proteins overflows the threshold of refolding, and cell death signaling is triggered in the ER.


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5.3 Proteasome Dysfunction Proteasomal dysfunction is also induced by aggregation of mutant proteins. As described in > Sect. 4, mutant polyglutamine proteins gather to the centrosome where proteasomal proteins are focused. The translocation by itself is rational, whereas elongated polyglutamine tract is hard to be cleaved by the proteasome (Venkatraman et al., 2004), and therefore the mutant peptide accumulates in the functional area of the proteasome. When large nonsoluble inclusion bodies are formed, the region is called ‘‘aggresome’’ (Johnston et al., 1998). At this stage, proteasomal function is disturbed by the aggregation.

5.4 Mitochondrial Stress Recent findings from many groups suggest that polyglutamine proteins induce mitochondrial stress. In addition to the results that polyglutamine proteins trigger cytochrome c release from mitochondria (Jana et al., 2001), it is reported that mutant htt proteins accumulate at mitochondrial membranes and they subsequently induce depolarization of mitochondrial membrane potential (Panov et al., 2002). Choo et al. (2004) reconfirmed that mutant htt protein associates with the outer‐mitochondrial membrane and lowers the Ca2þ threshold to open mitochondrial permeability transition pore.

5.5 Which Causes Symptoms, Neuronal Death or Neuronal Dysfunction? It is still controversial whether neuronal dysfunction or neuronal death causes the symptoms of polyglutamine disease seen in such patients. Recent results from transgenic mouse models have shown that symptomatic onset precedes neuronal death in the brain. Transgenic mice show motor dysfunction before the decrease in neurons is observed, suggesting strongly that neuronal dysfunction causes symptoms. In this sense, it is of note that mental retardation, which does not accompany neuronal death in youth, and polyglutamine diseases share common pathological molecules such as CBP and PQBP1. Both proteins are sequestered into polyglutamine inclusion bodies in neurons (Kazantsev et al., 1999; McCampbell et al., 2000; Steffan et al., 2000; Okazawa et al., 2002; Busch et al., 2003) to cause dysfunction of both molecules in transcription. At the same time, mutations of CBP and PQBP1 are the causes of Rubinstein–Taybi syndrome (Murata et al., 2001; Kalkhoven et al., 2003) and Renpenning syndrome (Kalscheuer et al., 2003; Lenski et al., 2003,), respectively. Therefore, disturbance of neuronal transcription may underlie the symptoms of neurodegenerative disorders before neuronal cell death occurs. Meanwhile, we do not know which kind of changes are induced in neurons after a long term of transcriptional dysfunction.

6

Frontier of Therapeutic Approaches

Significant advances have been made recently in the development of therapeutic approaches. Therefore, polyglutamine diseases are now being seen from being intractable to be curable. Various approaches proposed so far will be classified into three categories: (1) approaches for common pathologies of poly‐Q diseases, (2) approaches for specific pathologies in a poly‐Q disease, and (3) nonspecific approaches to inhibit neuronal death. The first category includes aggregation modifiers, cross‐linking inhibitors, transcriptional upregulators, caspase inhibitors, and RNAi. The second category includes antiandrogen drug for SBMA (Kennedy’s disease) and RNAi for disease‐specific SNPs. The third category includes various chemicals that have been investigated for other neurodegenerations. These candidates will be tested by cellular and animal (Drosophila, mouse, rat) models and then will be forwarded to human clinical trials. In the future, combination of different types of drugs will increase the efficiency of the therapy.

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6.1 Aggregation Modifiers It is still a focus of debate whether aggregation is toxic or protective for neurons. The classical idea is that aggregated mass of polyglutamine proteins is toxic for cells. However, Greenberg’s group and Zoghbi’s group reported that aggregation itself is not essential for cell toxicity (Klement et al., 1998; Saudou et al., 1998). They used cellular or animal models and tested the toxicity of mutant proteins lacking the aggregation domain or nuclear transport domain. Their data showed that nuclear transport but not aggregation induces cell death or dysfunction. Thereafter, numerous reports were published suggesting that nonaggregated form affects neuronal function or death. On the other hand, recent data have shown that intermediate forms during aggregation play critical roles in the pathology. These findings collectively suggest the idea that aggregation processes rather than final large aggregates are critical (Arrasate et al., 2004). According to progress in the research of pathophysiology, aggregation inhibitors are now tested. Wanker’s group reported that Congo red and antibodies are capable of inhibiting aggregation (Heiser et al., 2000). His group further screened chemicals by using their original automatic machine‐ quantifying aggregation in vitro and found that benzothiazole and some other chemicals are effective in suppressing aggregation (Heiser et al., 2002). Probably, as far as I know, the first approach to synthesize peptides that inhibit aggregation process was done by Thompson’s group. Her group constructed an expression vector for a polypeptide that contained two 25 repeats of glutamine intercalated by a‐helix sequence. This polypeptide binds to polyglutamine and inhibits aggregation (> Figure 12‐3). After checking

. Figure 12‐3 Therapeutic mechanism of aggregation inhibitor

the effect on aggregation in cell culture models, they proceeded to the Drosophila model and found that the peptide suppressed eye degeneration induced by htt (Kazantsev et al., 2002). Recently, Nukina’s group reported that trehalose suppresses aggregation and symptoms in vitro and in vivo (Tanaka et al., 2004).

6.2 Cross‐Linking Inhibition Aggregated proteins are cross‐linked by transglutaminase and become firm inclusion bodies. There is an idea that inhibition of cross‐linking will improve the pathology (Igarashi et al., 1998). Recently Karpuj et al.


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(2002) reevaluated this idea and reported that cystamine, an inhibitor of transglutaminase, effectively improved symptoms in model mice.

6.3 Cleavage Inhibition It is known that shorter polypeptides containing the polyglutamine tract are more toxic than full‐length proteins. This leads to the idea that inhibition of cleavage will be effective for therapy. Ona et al. (1999) applied ZVAD‐fmk, a caspase inhibitor to Huntington’s disease model mice and found that it repressed symptom progression and elongated survival. The group also showed that minocycline inhibits caspases and rescues Huntington’s disease mouse model.

6.4 Transcriptional Upregulation As described above, polyglutamine proteins interact with a number of nuclear proteins and disturb nuclear functions. Therefore, upregulation of transcription is considered to act against the pathology. HDAC inhibitors (SAHA, sodium butylate, etc.) have been shown to suppress symptoms and disease progression (McCampbell et al., 2001; Steffan et al., 2001; Hockley et al., 2003) (> Figure 12‐4). Since HDAC inhibitors have been used in clinics as anticancer drugs, clinical trials for polyglutamine diseases are already in progress. . Figure 12‐4 Therapeutic mechanism of HDAC inhibitors

6.5 RNAi RNAi is expected as the most direct method to suppress expression of mutant proteins. However, theoretically, some critical concerns are raised for this approach. RNAi suppresses not only mutant proteins but also normal proteins from normal allele genes. In addition, recent research has revealed that RNAi is not completely specific. RNAi suppresses not only the target gene but also unrelated genes to some extent. To overcome the first problem, Miller et al. used disease allele‐specific SNP sequence as the target of siRNA (> Figure 12‐5). They showed in the case of SCA3 that siRNA suppresses mutant poly‐Q protein specifically without affecting normal protein expression (Miller et al., 2003).

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. Figure 12‐5 Disease allele‐specific siRNA

6.6 Modification of Nuclear Transport Androgen receptor is a well‐known steroid hormone‐binding nuclear receptor. After binding to the ligand, the receptor moves to the nucleus and switches on transcription of target genes. Antiandrogen drugs have been developed for prostate hypertrophy and some relevant cancers. Antiandrogen drugs are classified into two groups, those suppressing nuclear transport of androgen receptor and those inhibiting receptor binding to the motif sequence of DNA. Sobue et al. reported that the former type, leuprorelin, remarkably suppresses symptoms of SBMA model mice (Katsuno et al., 2003).

6.7 Drugs for Mitochondria There are many candidate drugs that promote viability of neurons nonspecifically. Minocycline is an example that has attracted attention in the therapy of ALS. It possesses actions of mitochondria membrane stabilization and caspase inhibition (Chen et al., 2000). In Huntington’s disease, the drug is in phase II of clinical trials.

7

Conclusion

I reviewed the research progress of pathomechanisms and therapeutic trials of polyglutamine diseases. This chapter has explained common pathologies shared by different polyglutamine diseases, aggregation, and various cell stresses triggered during the aggregation process. The core pathology seems to be also shared by other neurodegenerations such as Alzheimer’s disease, Parkinson’s disease, tauopathy, and ALS. The pathology seems to be modified by neurons or neuron type‐specific cellular circumstances, although the details are not yet known sufficiently. Modification or suppression of these pathomechanisms will surely lead to the cure of these intractable diseases.

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The Proteasome, Protein Aggregation, and Neurodegeneration

S. Wilk

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373

2 2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.2 2.3 2.3.1 2.3.2 2.4 2.4.1 2.4.2 2.5 2.6 2.6.1 2.6.2 2.6.3 2.6.4 2.6.5 2.6.6 2.6.7 2.7

Biochemistry of the Proteasome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373 20S Proteasome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 Catalytic Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Immunoproteasome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 Proteasome Activators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 11S Regulator (PA28) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 PA200 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 26S Proteasome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 The 19S Regulatory Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Hybrid Proteasomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 The Ubiquitin‐Proteasome Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 380 Ubiquitin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 380 E1: Ubiquitin‐Activating Enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 380 E2: Ubiquitin‐Conjugating Enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 380 E3: Ubiquitin–Protein Ligase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 381 Targeting of Substrates for Ubiquitination and Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 381 E4: Ubiquitin Elongation Factor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 382 Ubiquitin Hydrolases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 382 Ubiquitin‐Independent Proteolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 382

3 3.1 3.1.1 3.1.2 3.1.3 3.2 3.2.1 3.3 3.4

Parkinson’s Disease and the Proteasome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 383 a‐Synuclein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 383 a‐Synuclein Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 384 a‐Synuclein and the Proteasome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 385 a‐Synuclein‐Interacting Protein: Synphilin‐1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386 Parkin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386 Search for the Parkin Substrate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 a‐Synuclein and Parkin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 Studies on the Proteasome in Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 389

4 4.1 4.2 4.2.1

Protein Aggregation and the Proteasome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 390 The Aggresome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391 Polyglutamine Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 Colocalization of Inclusions and Components of the Ubiquitin‐Proteasome System . . . . . . . . . . 393

#


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The proteasome, protein aggregation, and neurodegeneration

4.2.2 4.3 4.4 4.5 4.6

Interaction of Proteasome and Poly‐Q‐Containing proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 The Role of the Proteasome in Protein Aggregation and Inclusion Formation . . . . . . . . . . . . . . . . 395 Evidence That Aggregate Formation Is Responsible for Cellular Pathology . . . . . . . . . . . . . . . . . . . . 396 Evidence That Inclusions May Be Cytoprotective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397 Are Polyglutamine‐Expanded Proteins Proteasome Substrates or Proteasome Inhibitors? . . . . . 398

5

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399


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Abstract: The presence of intracellular inclusions containing aggregated proteins feature in affected neurons of patients with neurodegenerative disease. Such inclusions also contain immunoreactive ubiquitin and proteasome subunits. Similar perinuclear inclusions termed aggresomes are formed when cells are treated with proteasome inhibitors. Since unfolded proteins are predominantly degraded by the ubiquitinproteasome system (UPS), a dysfunction of this system has been proposed to contribute to neurodegeneration. This chapter presents an outline of the biochemistry of the UPS followed by an examination of the experimental evidence relating this protein-degrading system to Parkinson’s disease, to protein aggregation and aggresome formation, and to polyglutamine disease. List of Abbreviations: ARJP, autosomal recessive juvenile Parkinsonism; AZ, antizyme; CFTR, Cystic fibrosis transmembrane conductance regulator; CHIP, carboxyl terminus of Hsc-70-interacting protein; INF-g, Interferon gamma; NII, neuronal intranuclear inclusions; ODC, ornithine decarboxylase; Pael-R, Parkin-associated endothelin receptor-like receptor; PGPH, peptidyl glutamyl peptide hydrolyzing; PSI, Z-lle-Glu(O-tBu)-Ala-leucinal; UIM, ubiquitin-interacting motif; UPS, ubiquitin-proteasome system; Z, N-benzyloxycarbonyl

1

Introduction

In recent years there has been considerable interest in the role of abnormal proteolysis in general and a dysfunction of the ubiquitin‐proteasome system in particular in the etiology of neurodegenerative disorders. The presence of protein aggregates in inclusion bodies is a common feature of some neurodegenerative diseases. One of the hallmarks of Parkinson’s disease is the occurrence of cytoplasmic inclusions known as Lewy bodies in nigral dopaminergic neurons. Neuronal intranuclear inclusions (NII) are found in hereditary diseases such as Huntington’s disease and the spinocerebellar ataxias. The latter diseases are caused by the formation of aggregate‐prone proteins containing an expanded polyglutamine tract. Since the proteasome degrades misfolded proteins, a dysfunction of the ubiquitin‐proteasome system may lead to the accumulation and subsequent aggregation of these proteins. Both Lewy bodies and NII contain immunoreactive ubiquitin and proteasome components. When normal cells are treated with proteasome inhibitors, perinuclear centrosomal inclusions termed aggresomes form in a microtubule‐dependent manner. These inclusions also contain immunoreactive ubiquitin and proteasome components. It has therefore been proposed that Lewy bodies and NII may form because the capacity of the ubiquitin‐proteasome system is either impaired or overwhelmed. There is evidence that the inclusions serve as a site for recruitment of proteasome components, and that this redistribution may cause a depletion of the proteasome in other cellular compartments. It has also been proposed that aggregated proteins directly inhibit the proteasome by being trapped within its catalytic chamber. Mutations in the parkin gene underlie autosomal recessive juvenile Parkinsonism (ARJP). The identification of parkin as an E3 ubiquitin–protein ligase further strengthens the association between the ubiquitin‐proteasome system and neurodegenerative diseases. There is evidence for an underlying impairment of the ubiquitin‐proteasome system in brains of patients with Parkinson’s disease. However, there are many studies that question the role of the ubiquitin‐proteasome system in neurodegeneration and this research area is one of considerable controversy. The focus of this chapter is on the experimental evidence that either does or does not support a role of the ubiquitin‐ proteasome system in Parkinson’s disease and in the polyglutamine diseases. The chapter begins with an overview of the basic biochemistry of the ubiquitin‐proteasome system.

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Biochemistry of the Proteasome

The proteasome is a unique proteolytic complex that incorporates features not seen in classical proteinases. These include a very high molecular weight, a complex subunit structure, multiple catalytic activities residing on distinct subunits, a catalytic mechanism employing an N‐terminal threonine as nucleophile, and self‐ compartmentation. This ‘‘multicatalytic endopeptidase complex’’ (a term adopted by the Nomenclature

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Committee of International Union of Biochemistry) serves as the catalytic core of the ubiquitin‐proteasome system and is responsible for the bulk of intracellular extralysosomal proteolysis. The proteasome is a component of all eukaryotic cells and its function is necessary for cell growth and viability. The proteasome has been the focus of many investigations on the possible role of abnormal proteolysis in the etiology of neurodegenerative diseases.

2.1 20S Proteasome The 20S proteasome was first detected, purified, and biochemically characterized more than 25 years ago (Wilk and Orlowski, 1980; Orlowski and Wilk, 1981; Wilk and Orlowski, 1983). The early publications described the cleavage of the leucyl‐p‐nitroanilide bond of the chromogenic substrate N‐benzyloxycarbonyl‐ Gly‐Gly‐Leu‐p‐nitroanilide by the supernatant fraction of a bovine pituitary homogenate. The responsible enzyme was isolated and found to be a high‐molecular weight proteinase (700,000) composed of multiple subunits of molecular weights ranging from 24,000 to 30,000. Further studies on its specificity demonstrated that in addition to cleaving bonds after hydrophobic amino acids such as leucine, the enzyme could also catalyze hydrolysis after basic and acidic amino acids. Experiments with inhibitors and activators demonstrated that the three catalytic activities represented three distinct active sites residing in a ‘‘multicatalytic proteinase complex.’’

2.1.1 Structure The structure of the proteasome was determined by electron microscopy and by X‐ray crystallography. A primitive proteasome from Thermoplasma acidophilum contains two nonidentical subunits termed a and b. Each subunit forms a seven‐membered ring, and the proteasome is composed of four stacked subunit rings (Dahlmann et al., 1989). Immunoelectron microscopy localized the a subunits to the two outer rings and the b subunits to the two inner rings (Grziwa et al., 1991). The eukaryotic 20S proteasome has a structure similar to the proteasome from Thermoplasma acidophilum i.e., it is also composed of four rings of seven subunits each (Pu¨hler et al., 1992). Although the subunits of the eukaryotic proteasome are related to either the a‐ or b‐subunits of the Thermoplasma enzyme, the eukaryotic enzyme contains seven distinct a subunits and seven distinct b subunits (Zwickl et al., 1992). Its structure is that of a complex dimer i.e., a7b7b7a7 (Kopp, 1993). The primary sequences of all subunits from several species are known from the nucleotide sequences of recombinant cDNA clones (Tanaka, 1998). Homologies among the N‐terminal regions of subunits indicate that they are products of a related family of genes (Lee et al., 1990; Sorimachi et al., 1991). Although the proteasome is the major extralysosomal proteinase of the cell, the 20S proteasome by itself does not degrade native proteins and only very slowly degrades unfolded proteins. The physical basis for this phenomenon was revealed by the X‐ray crystallographic structure of the molecule. Solving the structure of the Thermoplasma enzyme at 3.4 A˚ located the active sites present on the b‐type subunits to the interior of this barrel‐shaped molecule. A narrow opening at each end of the particle provides a barrier against the entrance of native proteins (Lo¨we et al., 1995). Since the cellular concentration of the proteasome can exceed 1% of the soluble protein, this structure serves to protect the cell from uncontrolled proteolysis. The X‐ray crystallographic structure of the yeast enzyme solved to 2.4 A˚ resolution surprisingly shows that it is sealed at both ends by N‐terminal chains of a‐subunits. The only visible access of proteins to the catalytic interior of the molecule appears to be by very narrow side entrances (Groll et al., 1997). The proteasome structure has been termed by Baumeister as a ‘‘self‐compartmentalizing protease’’ (Baumeister et al., 1998). Efficient protein and peptide degradation is assured by the combination of the 20S proteasome with cellular regulatory proteins. The sealed form of the proteasome is described in the literature as a ‘‘latent’’ form. The latent form can be converted to an open channel form during the purification procedure or by low concentrations of SDS, some long‐chain fatty acids, cardiolipin, or hydrophobic peptides (Wilk and Orlowski, 1983; Dahlmann et al., 1985; Ruiz de Mena, 1993; Wilk and Chen, 1997).


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2.1.2 Catalytic Activities Initial characterization of the bovine pituitary proteasome identified three distinct catalytic activities that cleaved peptide bonds after hydrophobic, basic, and acidic amino acids. These were termed chymotrypsin‐ like‐, trypsin‐like‐, and peptidyl glutamyl peptide hydrolyzing (PGPH) activities, respectively. Only three of the seven distinct b‐type subunits of the eukaryotic proteasome are known to be catalytically active. A study of wild‐type‐ and mutant yeast proteasomes assigned each of the catalytic activities to a given b subunit. The b‐1 subunit is responsible for the PGPH activity, the b‐2 subunit for the trypsin‐like activity, and the b‐5 subunit for the chymotrypsin‐like activity (Dick et al., 1998). The function of the remaining four b subunits remains unknown. The possibility that the mammalian proteasome contains more than three active sites is not excluded. The bovine liver enzyme crystal structure was determined at 2.75 A˚ resolution. The yeast and liver enzymes have the same subunit arrangement. However, on the basis of localization of functional groups, Unno et al. (2002) proposed a catalytic role for the b‐7 subunit. They found that the cleft for this putative active site was shallower than the pockets of the b‐1, ‐2 and ‐5 subunits suggesting that a small neutral amino acid could bind in the P1 position. This would fit with the proposal of a distinct small neutral amino acid preferring activity described by Orlowski et al. (1993). A detailed discussion of the catalytic activities of the proteasome can be found in the review of Orlowski and Wilk (2000).

2.1.3 Mechanism Proteolytic enzymes have been traditionally divided into four classes representing their catalytic mechanism i.e., serine‐, cysteine‐, metallo‐, and aspartyl proteases. The proteasome is a member of a new mechanistic class and employs an N‐terminal threonine as the active site nucleophile. Evidence in support of this assignment comes from several different studies (reviewed in Orlowski and Wilk, 2000). Lactacystin, a selective inhibitor of the proteasome, forms a covalent bond with the N‐terminal threonine of the b‐5 subunit (Fenteany et al., 1995). Mutation of the N‐terminal threonine residue of the b subunit of the Thermoplasma proteasome to alanine, or its deletion, abolishes enzymatic activity (Seemuller, 1995). Finally support for this assignment is derived from crystallographic analysis of the proteasome from Saccharomyces cerevisiae in complex with an inhibitor (Groll, 1997). The proteasome is a member of the newly recognized class of N‐terminal nucleophile hydrolases (Brannigan et al., 1995; Duggleby et al., 1995).

2.1.4 Inhibitors Proteasome inhibitors have been pivotal in uncovering the physiological significance of this proteinase complex. Early studies on the proteasome established the utility of peptidyl aldehydes as proteasome inhibitors (Wilk and Orlowski, 1983). Peptidyl aldehydes inhibit serine, cysteine, and threonine proteinases by reacting with the active site nucleophile to form a hemiacetal (or in the case of cysteine proteinases a thiohemiacetal adduct). Lack of specificity is a potential limitation of these compounds. The trypsin‐like component of the proteasome is inhibited by the peptidyl aldehyde leupeptin, but this compound inhibits many enzymes including the calpains and some cathepsins. The first inhibitor of the chymotrypsin‐like activity N‐benzyloxycarbonyl (Z)‐Gly‐Gly‐leucinal synthesized as an analog of the substrate Z‐Gly‐Gly‐Leu‐ p‐nitroanilide, is not very potent. In 1994, the peptidyl aldehydes Z‐Ile‐Glu(O‐tBu)‐Ala‐leucinal (PSI) (Traenckner et al., 1994) and Z‐Leu‐Leu‐Leucinal (MG132) (Palombella et al., 1994) were used to demonstrate the role of the proteasome in the activation of the transcription factor NF‐kB. Subsequently proteasome inhibitors have been established as essential tools in the study of proteasome function. The Streptomyces metabolite lactacystin is a proteasome inhibitor of greater specificity than the aldehydes (Fenteany et al., 1995), but also inhibits cathepsin A (Ostrowska et al., 1997) and tripeptidyl peptidase II (Geier et al., 1999). Many different classes of reversible and irreversible proteasome inhibitors have been developed (for review, see Myung et al., 2001). Of note is bortezomib, a potent and specific dipeptidyl boronate, approved for the treatment of multiple myeloma (Adams, 2003).

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2.2 Immunoproteasome In higher eukaryotes, the three catalytically active subunits can be replaced by subunits whose expression is upregulated by the cytokine interferon (IFN)‐ g. The b‐1 subunit (Y) is replaced by LMP2 (b1i), the b‐2 subunit (Z) by LMP10 (b2i, MECL‐1), and the b‐5 subunit (X) by LMP7 (b5i). This form of the proteasome has been designated as ‘‘immunoproteasome’’ to reflect its probable role in the generation of antigenic peptides for presentation by the MHC class I system (Tanaka and Kasahara, 1998). As a result of the subunit replacements, there is a change in the specificity of the proteasome such that cleavage of peptide bonds after hydrophobic and basic amino acids is favored, and cleavage after acidic amino acids is suppressed (Aki et al., 1994). Although many tissues contain mixed populations of proteasomes, the pituitary almost exclusively contains the constitutive form, whereas the spleen almost exclusively contains the immunoproteasome (Eleuteri et al., 1997; Orlowski et al., 1997). The PGPH activity is almost totally absent in the purified spleen proteasome (Eluteri et al., 1997). The LMP2 subunit replaces the b‐1 subunit and expresses a chymotrypsin‐like activity (Orlowski et al., 1997).

2.3 Proteasome Activators 2.3.1 11S Regulator (PA28) A protein activator of the 20S proteasome termed 11S regulator (REG) or PA28 was independently described by two groups (Dubiel et al., 1992a; Ma et al., 1992). The activator is composed of two homologous 30‐kDa subunits designated a and b (Mott et al., 1994). Most recent evidence indicates that the activator is a heptamer, although the exact arrangement of subunits is unclear (Zhang et al., 1999). The crystal structure of the homoheptameric recombinant a subunit has been solved (Knowlton et al., 1997). Combination of 11S REG with the proteasome dramatically stimulates the degradation of small synthetic peptide substrates but not the degradation of proteins (Dubiel et al., 1992b; Ma et al., 1992). This is due both to an increase in the Vmax and a decrease in the Km of the substrate. Thus, 11S REG behaves as a positive allosteric activator (Ma et al., 1992). Since the expression of both a‐ and b genes is strongly upregulated by IFN‐g the activator has been proposed to play a role in the processing of peptides that associate with MHC class I molecules (Realini et al., 1994). Insight into the mechanism of activation was gained by crystallization of the 11S REG from Trypanosoma brucei with the proteasome from yeast (Whitby et al., 2000). Binding of the activator induces conformational changes in the N‐terminal tails of a subunits causing an opening of the gate leading to the interior of the catalytic chamber. Cloning of the activator subunits led to the identification through a database search of a homologous protein of unknown function named Ki antigen, previously identified in the sera of patients with systemic lupus erythematosus (Nikaido et al., 1990). Because of its similarity to a and b subunits of the activator, its copurification with the activator and its immunoprecipitation by anti‐20S proteasome antibodies, it has been renamed as PA28g or REGg (Tanahashi et al., 1997). It is a homoheptamer and also a proteasome activator (Wilk et al., 2000). The cellular distribution of REGg is distinct from that of REGab. REGg is primarily found in the nucleus but not the nucleolus, whereas REGab is found primarily in the cytoplasm and also in the nucleolus (Wojcik et al., 1998). Disruption of the REGg gene in mice impairs body growth and slows the growth of mouse embryonic fibroblasts and impedes their entry into S phase (Murata et al., 1999). The physiological significance of this nuclear proteasome activator still remains unknown. REGg has been reported either as a selective activator of the trypsin‐like activity (Li et al., 2001) or a general activator of the three catalytic activities of the proteasome (Wilk et al., 2000). These apparent discrepancies were resolved when it was shown that the properties of REGg were altered by the method of purification (Gao et al., 2004). Thus, treatment with ammonium sulfate converts the form that selectively stimulates the trypsin‐like activity to a general activator. It is of interest that this switch can also be achieved by mutagenesis of Lys188 to Asp or Glu (Li et al., 2001). The physiological significance of the alteration in properties of REGg remains to be determined.


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2.3.2 PA200 The newest of the proteasome activators to be described is PA200 (Ustrell et al., 2002). This 200‐kDa nuclear protein purified from bovine testis resembles the REG (PA28) family in that it stimulates the hydrolysis of small peptide substrates in an ATP‐independent manner but does not stimulate the hydrolysis of proteins. All three catalytic activities of the proteasome are stimulated by PA200 with the greatest effect exerted on the PGPH activity. In general, activation is less robust that that produced by the REG family. Western blot analysis shows that PA200 is most abundant in testis whereas other tissues contain a 160‐kDa species. Of interest is the staining of a prominent 60‐kDa band from brain samples. PA200 is subject to posttranslational modification but it is unclear how such modification affects its properties. PA200 contains repeat motifs that resemble ARM/HEAT repeats (Kajava et al., 2004). They serve to impart a solenoid structure to PA200 and may facilitate binding to the 20S proteasome. Electron microscopic analysis of PA200–20S complexes show singly and doubly capped forms (Ortega et al., 2005). PA200 appears to activate the proteasome by opening the axial gate of the a‐rings and unlike the REG family, binds to the proteasome as a monomer. The physiological significance of PA200 is obscure but it has been proposed to play a role in DNA repair (Ustrell et al., 2002).

2.4 26S Proteasome A high‐molecular weight ubiquitin–lysozyme conjugate degrading enzyme sedimenting at 26S was partially purified from reticulocyte lysates in 1986 (Hough et al., 1986). This enzyme was purified by two groups in 1987 (Hough et al.,1987; Waxman et al., 1987). The 26S enzyme largely precipitates at 0–38% ammonium sulfate saturation whereas 40–70% saturation is required to precipitate the 20S proteasome (Waxman et al., 1987). Hough et al. (1987) purified two high‐molecular weight proteolytic enzymes from a rabbit reticulocyte lysate using chromatography and glycerol gradient centrifugation. The smaller of the two enzymes was identified as the 20S proteasome on the basis of its appearance on SDS‐PAGE gels. The higher molecular weight 26S enzyme degraded 125I–lysozyme–ubiquitin conjugates in an ATP‐dependent manner. SDS‐PAGE analysis showed the presence of bands corresponding to the 20S proteasome along with a characteristic set of bands ranging from 34 kDa to 110 kDa. A highly purified preparation of the 26S proteasome from rat liver clearly showed that it was composed of a core 20S proteasome with multicatalytic properties and an additional structure composed of proteins from 35 kDa to 110 kDa, presumably serving a regulatory role (Ugai et al., 1993).

2.4.1 Structure The true sedimentation coefficient of the 26S proteasome is actually 30S. Although it was proposed to rename this complex the 30S proteasome (Yoshimura et al., 1993), the 26S name has persisted. Its molecular weight is approximately 2.5 million. Electron microscopy revealed a ‘‘caterpillar’’ shape and a symmetrical assembly of two domains (Walz et al., 1998). The thinner central domain contains four protein layers that are the rings of the 20S proteasome. The regulatory complexes attached to each end of the core particle assume a U‐shape and are composed of about 18 proteins.

2.4.2 The 19S Regulatory Complex A multiprotein complex referred to as the m particle was purified from embryos of Drosophila (Udvardy, 1993). This protein complex possessed no proteolytic activity but was incorporated in an ATP‐dependent manner into a larger complex that displayed strong proteolytic activity. The larger complex is now known to

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be the 26S proteasome and the m particle is the 19S regulatory complex (also termed 19S cap complex (Peters et al., 1994)) or PA 700 (DeMartino et al., 1994). Sequencing of one of the regulatory subunits identified it to be a member of an ATPase family (Dubiel et al., 1992b). This family is termed AAA for ATPase Associated to a variety of cellular Activities (Confalonieri and Duguet, 1995). The regulatory complex contains a ring of six AAA ATPases that binds to the outer a subunits of the 20S proteasome (Glickman et al., 1998b). The ATPases presumably provide energy for protein unfolding, opening of the entrance gate to the catalytic chamber of the proteasome, and translocation of the substrate. These subunits in the yeast regulatory particle have been termed Rpts for regulatory particle triple A proteins (Finley et al., 1998). The remainder of the regulatory particle subunits are termed Rpns for regulatory particle non‐ATPases. Mutation of one of the ATPases, Rpt2, decreases the peptidase activity of the proteasome possibly by affecting the gating of the channel (Rubin et al., 1998). When the Rpn10 subunit is deleted in yeast, a subcomplex containing eight non‐ATPase subunits, termed the lid, is dissociated from the yeast 26S proteasome (Glickman et al., 1998a). The lid represents the distal portion of the regulatory complex. The remaining regulatory subunits that are attached to the core 20S proteasome comprise the base. The Rpn10 deletion mutant lacking the lid is capable of degrading nonubiquitinated proteins but not ubiquitin–protein conjugates. The base contains the six ATPases plus two non‐ATPase subunits Rpn1 and Rpn2. It has ATP‐dependent chaperone activity and a binding site for unfolded proteins (Braun et al., 1999). This may allow the base to discriminate between folded and unfolded states of the protein. The base also appears to contain the subunit responsible for the recognition of polyubiquitin chains. When ubiquitin was mutated to introduce a cysteine residue and the tetraubiquitin form of the mutant ubiquitin was cross‐linked to the purified bovine 26S proteasome, an ATPase subunit Rpt5 (S60 ) was identified as the target of binding (Lam et al., 2002). Earlier studies had pointed to subunit Rpn10 (S5a) as the target of binding (Deveraux et al., 1994). However, disruption of the Rpn10 gene in yeast results in only a mild phenotype (van Nocker et al., 1996). It does however remain possible that Rpn10 is responsible for the recognition of a subset of ubiquitinated substrates. Elsasser et al. (2002) have shown that Rpn10 can contribute to the binding of ubiquitin chains by the intact proteasome. A tetraubiquitin chain appears to be the minimum signal for recognition by the 26S proteasome (Piotrowski et al., 1997). Ubiquitin–protein conjugates must first be deubiquitinated and unfolded before they can enter the 20S proteasome. Evidence has been presented that the lid subunit Rpn11 is responsible for deubiquitination and that surprisingly this subunit appears to be a metalloproteinase (Verma et al., 2002; Yao and Cohen, 2002). Although classical deubiquitinating enzymes are cysteine proteinases and are inhibited by ubiquitin aldehyde, deubiquitination by the 26S proteasome proceeds in the presence of ubiquitin aldehyde but is sensitive to inhibitors of metalloenzymes. Among the non‐ATPases, Rpn11 is the most conserved subunit. This subunit as well as a subunit of the COP signalosome contains a distinct arrangement of histidines, aspartate, and glutamate, EXnHXHX10D, highly suggestive of a metalloproteinase. Mutation of the putative active site histidines is lethal in yeast and leads to stabilization of ubiquitin–protein conjugates. It is proposed that a major function of the lid is to serve as a specialized isopeptidase, coupling deubiquitination to degradation. A distinct isopeptidase present as an integral component of the 19S regulatory complex was also described (Lam et al., 1997). This enzyme is sensitive to ubiquitin aldehyde and removes one ubiquitin at a time from the distal end of a chain. It was proposed to play an editing role by removing ubiquitin from poorly ubiquitinated substrates, thereby freeing them from degradation by the proteasome. Subunits of the lid contain structural motifs also present in the COP9/signalosome complex and in eIF3, a mediator of translational initiation (Glickman et al., 1998a).

2.5 Hybrid Proteasomes The term hybrid proteasome designates a form in which the 20S proteasome is capped at one end by a 19S regulatory subunit complex and at the other end by the 11S REG (PA28) (Tanahashi et al., 2000). It is notable that the content of hybrid proteasome in HeLa cells was estimated to be equivalent to that of the 26S


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proteasome (Tanahashi et al., 2000). Hybrid proteasomes can be reconstituted in vitro from purified 20S proteasome, 19S regulatory complex, and 11S REG (PA28) (Kopp et al., 2001; Cascio et al., 2002). It is notable that the 11S REG–20S proteasome complex does not degrade proteins. The hybrid proteasome possessing a 19S regulatory complex cap can degrade ornithine decarboxylase in an ATP‐dependent manner (Tanahashi et al., 2000). The hybrid proteasome may serve a special role in the immune system by directly degrading proteins to antigenic peptides. Presumably the protein will be recognized by and unfolded by the19S regulator cap. Binding of the 11S REG at the other end will facilitate the exit of the peptide products by opening the exit port. It was reported that the pattern of peptide products generated by the 26S proteasome and the hybrid proteasome differ (Cascio et al., 2002). A hybrid proteasome has been described in S. cerevisiae in which the 20S proteasome is capped at one end by the 19S regulatory particle and the other end by Blm10, the yeast ortholog of PA200 (Schmidt et al., 2005) (> Figure 13-1).

. Figure 13-1 Proteasome forms: The cell contains a heterogeneous population of proteasomes consisting of free 20S proteasome and 20S proteasome capped with protein regulators (Tanahashi et al., 2000). Not shown: singly capped forms and PA200‐containing hybrid proteasomes

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2.6 The Ubiquitin‐Proteasome Pathway 2.6.1 Ubiquitin The discovery of the ubiquitin system for protein degradation stemmed from investigations into the unexpected observation that intracellular protein degradation requires ATP. Early studies by Hershko, Ciechanover and colleagues utilized reticulocyte lysates as a cell‐free system for the study of ATP‐dependent proteolysis. By simple fractionation over a DEAE–cellulose column, they were able to separate a small heat‐ stable protein termed ATP‐dependent proteolysis factor (APF‐1) from the effluent. APF‐1, while having no proteolytic activity itself, stimulated ATP‐dependent protein breakdown when added to the column eluate (Hershko et al., 1979). APF‐1 was identified as ubiquitin, a highly conserved and widely distributed 76‐amino‐acid polypeptide (Wilkinson et al., 1980). Ubiquitin was known to form a covalent bond through its C‐terminal to the E‐amino group of a lysine residue of histone 2A. Histone 2A is monoubiquitinated. For other proteins, the bound ubiquitin in turn could donate a lysine residue for isopeptide bond formation with a second ubiquitin molecule. By continuing this process, a polyubiquitin chain can be assembled. The minimum signal for recognition by the proteasome is a tetraubiquitin chain (Thrower et al., 2000). For an in‐depth and comprehensive review of the ubiquitin‐proteasome system the reader is referred to Glickman and Ciechanover (2002).

2.6.2 E1: Ubiquitin‐Activating Enzyme The ubiquitin‐activating enzyme E1, a homodimer, was purified by ubiquitin affinity chromatography (Ciechanover et al., 1982). The enzyme activates ubiquitin by the formation of a thiol ester in a reaction that utilizes ATP and also forms AMP þ PPi. The ATP‐dependent reaction of E1 with ubiquitin can be summarized as: Ubiquitin þ ATP þ E1 SH ! E1 S ubiquitin þ AMP þ PPi : There is a single ubiquitin‐activating enzyme in the yeast genome termed UBA1, and this gene is essential for viability. The ts85 mouse mammalian carcinoma cell line harbors a temperature sensitive mutation in E1 (Finley et al., 1984). When these cells are brought to the nonpermissive temperature, there is a dramatic reduction in the degradation of short‐lived proteins. Therefore ubiquitin conjugation is necessary for most of the degradation of short‐lived proteins in vivo (Ciechanover et al., 1984). E1 by itself is not sufficient to catalyze the formation of ubiquitin–protein conjugates. In an extension of the earlier affinity chromatography procedure for purifying E1, two additional enzymes were isolated (Hershko et al., 1983). One of these termed E2 was eluted with DTT, and required both ATP and E1 for binding to the column. The other enzyme termed E3 did not require ATP for binding and was eluted with salt or high pH. All three enzymes were required for the efficient conjugation of ubiquitin to proteins.

2.6.3 E2: Ubiquitin‐Conjugating Enzyme The ubiquitin‐conjugating enzyme E2 serves as an acceptor of the ubiquitin moiety from E1 by also forming a thiol ester. The reaction is: E1 S ubiquitin þ E2 SH ! E1 SH þ E2 S ubiquitin: Ubiquitin conjugating enzymes are termed Ubcs and represent a large family of enzymes. There are eleven Ubcs in the yeast genome plus two additional family members that serve to conjugate ubiquitin‐like proteins. Higher eukaryotes have many more Ubcs. Ubcs can directly transfer ubiquitin to a target protein or more commonly transfer ubiquitin to a protein that is associated with an E3. The Huntington’s disease


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gene product huntingtin interacts with E2–25K (Kalchman et al., 1996). This may be an example in which an E2 transfers its ubiquitin to the target protein directly rather than to a target protein bound to an E3. E2–25K is highly expressed in brain and has strong similarity to yeast UBCs 1, 4, and 5. These UBCs play a role in degrading abnormal proteins.

2.6.4 E3: Ubiquitin–Protein Ligase E3s play a central role in targeting the protein for ubiquitination. E3‐bound proteins are ubiquitinated by the E2–ubiquitin complex. The reaction is: E2 ubiquitin þ E3 protein ! Ubiquitin protein conjugate þ E2 þ E3: E3s represent a diverse group of proteins or protein complexes. The two major classes of ubiquitin protein ligases are the HECT domain E3s and the RING finger E3s. Members of the HECT domain family have a 350‐amino‐acid sequence that is homologous to the E6‐ associated protein carboxyl terminus. E6‐associated protein (E6‐AP) was the first recognized member of this family and targets p53 for degradation (Huibregtse et al., 1993). The HECT domain proteins contain a conserved Cys that receives ubiquitin transferred from an E2. It is of interest that mutations in the E6‐AP gene cause Angleman’s syndrome, a severe form of mental retardation (Kishino et al., 1997). This points to an abnormality in the degradation of brain proteins by the ubiquitin‐proteasome system in this disease. The amino‐terminal domains of these proteins vary suggesting that they may play a role in substrate recognition. RING finger motifs consist of a cross brace structure of conserved cysteine and histidine residues that bind two zinc cations. The ring finger domain likely binds E2. Some ring finger E3s are monomeric or homodimeric proteins. One such E3 is parkin (see > Sect. 3.2). Other E3s are part of a multisubunit complex. Examples of the latter are the anaphase‐promoting complex (APC) or cyclosome, involved in cell cycle regulation and which is composed of at least 11 subunits, the skp‐1‐cullin/Cdc53‐F‐box proteins (SCF), and the von Hippel Lindau Elongin BC (VBC) cullin/skp‐1/F‐box protein complex. The F‐box protein of the SCF complex is variable and appears to govern recognition of the substrate protein. The SCF complexes primarily target phosphorylated proteins. The VBC complex targets hypoxia‐inducible factor 1 for degradation.

2.6.5 Targeting of Substrates for Ubiquitination and Degradation The targeting of a protein to the ubiquitin‐proteasome system can be achieved in multiple ways. These mechanisms are as yet incompletely understood and are the focus of intense investigation (for a detailed review see Glickman and Ciechanover, 2002). Some proteins such as IkB‐a require prior phosphorylation (Traenckner et al., 1994) and the SCF class of E3s recognizes phosphorylated proteins. Mitotic cyclins contain a conserved nine‐amino‐acid sequence termed the destruction box (Yamano et al., 1998). This sequence may serve as a recognition motif for E3, known as the anaphase promoting complex or cyclosome (Sudakin et al., 1995). In some cases, E3 recognizes an ancillary protein. This is seen in E6‐AP, the first member of the HECT class of E3s in which the human papilloma virus protein E6 binds both the substrate p53 and E3 (Scheffner et al., 1994). The resulting ternary complex formation is necessary for ubiquitination and degradation. Some substrates obey the N‐end rule i.e., the nature of the N‐terminal amino acid governs the stability of the protein (Varshavsky, 2000). Substrates may be also be targeted by exposure of a hydrophobic surface. This appears to be the case for the degradation of the yeast mating type transcription factor a2 (Laney and Hochstrasser, 1999). It is proposed that uncovering of a normally buried hydrophobic surface may serve as a general recognition signal for the degradation of damaged proteins. Finally, sequences rich in proline, glutamate, serine, and threonine (PEST sequences) also destabilize proteins (Rechsteiner and Rogers, 1996).

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2.6.6 E4: Ubiquitin Elongation Factor A factor in yeast that could elongate ubiquitin chains was isolated and termed E4 (Koegl et al., 1999). E4 binds to ubiquitin moieties of some short ubiquitin‐chain‐conjugated proteins in concert with E1, E2, and E3. Since the minimum signal for recognition by the proteasome is a tetraubiquitin chain, the action of E4 ensures efficient polyubiquitination and proteolysis. The E4 of yeast is a protein known as UFD2. The structure of UFD2 is highly conserved with strong homology within its C‐terminal domain. This domain is termed the U box for UFD‐2 homology domain. UFD2 is not essential for viability in yeast but appears to be important under conditions of cell stress. Carboxyl terminus of Hsc‐70‐interacting protein (CHIP) contains a U‐box motif and has E3 ligase activity. It may play a role in facilitating the E3 activity of parkin (see > Sect. 3.2). The U‐box protein E4B binds to and accelerates the degradation of ubiquitinated ataxin‐3 (see ‘‘Ataxin‐3’’).

2.6.7 Ubiquitin Hydrolases The posttranslational modification of ubiquitination is a reversible process. Ubiquitin hydrolases disassemble polyubiquitin chains that are ligated to proteins via isopeptide bond linkages and also serve to cleave the head‐to‐tail fusions of polyubiquitin chains that are the products of the ubiquitin gene. The isopeptide bonds of the protein‐linked ubiquitin chains are formed by coupling the carboxyl group of the terminal glycine of the proximal residue to the E‐amino group of lys48 of the distal residue, although linkage to lys63 has also been observed. Ubiquitin hydrolases are of importance in the recycling of ubiquitin, maintenance of appropriate cellular levels of free ubiquitin, and in governing the stability of ubiquitin–protein conjugates. Two major families of deubiquitinating enzymes have been described (for review see Wilkinson and Hochstrasser, 1998). Members of these families are cysteine proteases. The first family, termed ubiquitin C‐terminal hydrolases (UCH), is related to PGP 9.5 (UCHL‐1), a highly abundant neuron‐specific protein. It is of interest that a missense mutation in the fourth exon of the UCHL‐1 gene has been identified in a German family with Parkinson’s disease (Leroy et al., 1998) and that UCHL‐1 immunoreactivity is found in Lewy bodies (Lowe et al., 1990). UCHs disassemble ubiquitin fusion proteins. The second family is referred to as ubiquitin‐specific‐processing proteases (UBPs). The UBP family represents a large and divergent class. There are at least 17 genes encoding putative UBPs in yeast. UBPs disassemble ubiquitin–protein conjugates by cleaving the Ub–Ub isopeptide bond. As discussed in > Sect. 2.4.2, subunit Rpn11 of the lid is responsible for deubiquitination and this subunit surprisingly appears to be a metalloproteinase.

2.7 Ubiquitin‐Independent Proteolysis Some proteins are degraded by the proteasome in an ubiquitin‐independent manner (for review see Orlowski and Wilk, 2003). An established ubiquitin‐independent substrate of the 26S proteasome is ornithine decarboxylase (ODC). Its degradation is dependent on the presence of ATP and antizyme (AZ) (Bercovich et al., 1989; Murakami et al., 1992). A C‐terminal domain of AZ promotes its binding to ODC, and an N‐terminal domain stimulates ODC degradation by the proteasome. Other proteins that are degraded by the 26S proteasome in an ubiquitin‐independent manner include: c‐Jun (Jarel‐Encontre et al., 1995), calmodulin (Tarcsa et al., 2000), the cyclin‐dependent kinase inhibitor p21Cip1 (Sheaff et al., 2000), and the tumor suppressor protein p53 (Asher et al., 2002). The free 20S proteasome is present in cells in relatively high concentrations. Its concentration in HeLa cells is about twice as high as the amount of the 26S or 20S–11S–REG complexes (Tanahashi et al., 2000). In general, natural unfolded proteins and oxygen‐damaged, misfolded, mutated, or damaged proteins are susceptible to degradation by the 20S proteasome. Mechanisms that lead to protein targeting, protein unfolding, and opening of the gated channel to the catalytic sites independent of ubiquitin and the 19S regulatory complex must exist for the 20S proteasome to degrade proteins. Targeting of a substrate to the


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proteasome could be brought about by an accessory molecule or by a sequence within the substrate. Similarly, opening of the gated channel could be effected by either an accessory molecule or by the substrate itself. This mechanism may explain the observation that low micromolar concentrations of the arginine‐ rich histone H3 accelerates the degradation of casein and lysozyme (Orlowski, 2001). There is direct evidence that some substrates themselves promote gating of the proteasome. The unfolded proteins p21Cip1 and a‐synuclein are as efficiently degraded by the closed‐channel (latent) form of the 20S proteasome as by the fully assembled activated 26S form (Liu et al., 2003). The 20S proteasome degrades oxidized proteins and increased surface hydrophobicity was proposed as a mechanism facilitating this process (Davies, 2001). It is of interest that hydrophobic peptides stimulate the degradation of synthetic substrates (but not proteins) by the 20S proteasome (Wilk and Chen, 1997). The 26S proteasome is not as effective in catalyzing the degradation of oxidized proteins even in the presence of ATP and ubiquitin (Grune et al., 1997). In vivo substrates of the 20S proteasome relevant to neurodegenerative diseases include a‐synuclein, an unfolded brain protein in which mutations have been discovered in Parkinson’s disease (Tofaris et al., 2001), and the microtubule‐associated protein tau (Cardozo et al., 2002; David et al., 2002).

3

Parkinson’s Disease and the Proteasome

Parkinson’s disease, a relatively common neurodegenerative disorder characterized by rigidity, bradykinesia, and resting tremor is associated with a progressive loss of dopaminergic neurons in the pars compacta of the substantia nigra. A defining pathological feature of idiopathic Parkinson’s disease is the presence of ubiquitin‐containing cytoplasmic inclusions known as Lewy bodies in affected dopaminergic neurons. Although the etiology of Parkinson’s disease is unknown, recent findings have suggested that a dysfunction of the ubiquitin‐proteasome system may play a contributing role. Genetic studies have led to the identification of several proteins that may play a role in the pathology of early onset forms of Parkinson’s disease. The relationship of these proteins to the ubiquitin‐proteasome system has been the subject of intense investigation.

3.1 a‐Synuclein In 1993, Ueda et al. detected a previously unrecognized constituent of an amyloid preparation that was not the amyloid‐b protein. The full‐length cDNA of the precursor of this component encoding a 140‐amino‐ acid protein was isolated. The protein was strongly hydrophobic and secondary structure predictions indicated that it has a strong tendency to form a b‐sheet configuration. Because of its tendency to form b‐structures, it was proposed as a factor promoting the process of amyloid formation perhaps by acting as a seed or core. The following year, two related proteins were purified and sequenced from human brain (Jakes et al., 1994). These were termed a‐ and b‐synucleins because of their similarity to a protein from Torpedo californica termed synuclein (Maroteaux et al., 1988). a‐Synuclein was found to be identical to the nonamyloid‐b protein described by Ueda et al. (1993). a‐Synuclein is an abundant brain protein of unknown function. Its concentration at the nerve terminal suggests a role in neurotransmission. a‐synuclein/ mice are viable and do not display any gross pathological changes (Abeliovich et al., 2000) and a‐synuclein is not required for neuronal development and differentiation. However, the electrically evoked release of dopamine is altered in these animals and the striatal dopamine content is significantly reduced. a‐Synuclein may therefore play a role in dopaminergic neurotransmission. Spillantini et al. (1997) showed that Lewy bodies from patients with idiopathic Parkinson’s disease stained strongly with antibodies against a‐synuclein. Lewy bodies from patients with dementia with Lewy bodies also were immunopositive for a‐synuclein. It was suggested that aggregation of a‐synuclein may underlie Lewy body formation in Parkinson’s disease and that the Lewy body may be central to the neurodegenerative process. Genetic markers on human chromosome 4q21‐q23 segregate with the Parkinson’s disease phenotype in a family of Italian descent. The human a‐synuclein gene was previously shown to map to this region. Therefore the location of the a‐synuclein gene established it as an excellent candidate gene for Parkinson’s

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disease. Genotype analysis revealed a single base change in the fourth exon of the a‐synuclein gene resulting in an A53T mutation, which segregated with the Parkinson’s disease phenotype (Polymeropoulos et al., 1997). The substitution of Thr for Ala is predicted to extend the b‐sheet structure, thereby favoring aggregation. Conway et al. (1998) found that although the structures of the wild type and the A53T mutant forms of a‐synuclein were indistinguishable, the mutant form showed a much greater propensity to fiberize. Although it is not likely that this mutation accounts for the majority of sporadic or familial cases of Parkinson’s disease, it may play a significant role in the early onset form of this disorder by facilitating the formation of Lewy bodies. A second mutation of A30P was identified in a German family (Kruger et al., 1998). The consequences of expressing mutant A53T or wild‐type a‐synucleins in transgenic mice was explored by Lee et al. (2002). Expression of human a‐synuclein harboring an A53T mutation in transgenic mice led to a late onset neurodegenerative disease whereas transgenic mice expressing high levels of wild‐ type a‐synuclein or the A30P mutation did not develop pathology. Transgenic mice expressing human a‐synuclein were generated by Masliah et al. (2000) who observed that by 2 months of age the mice developed intraneuronal inclusions in the nucleus and cytoplasm, which were immunoreactive to a‐synuclein. Moreover tyrosine hydroxylase‐positive neurons within the substantia nigra contained abnormal accumulations of a‐synuclein and immunoreactive ubiquitin. Although the inclusions had some similarities to Lewy bodies, they differed by their presence in the nucleus and their lack of fibrillar components. The mice also displayed deficits in motor performance. The authors suggested that increased expression or accumulation of a‐synuclein could play an important role in the pathogenesis of Lewy body neurodegenerative disease.

3.1.1 a‐Synuclein Degradation a‐Synuclein is one of the few well‐documented examples of a protein that can be degraded by the proteasome in an ubiquitin‐independent manner. Its native unfolded structure makes it susceptible to degradation by the 20S proteasome (Orlowski and Wilk, 2003). Tofaris et al. (2001) prepared stable transfectants of SH‐SY5Y cells containing overexpressed a‐synuclein. When these cells were treated with lactacystin, levels of nonubiquitinated a‐synuclein increased. Incubation of purified recombinant a‐synuclein with 20S proteasome confirmed direct degradation. The degradation of wild type and mutant A53T a‐synuclein was studied in the same neuroblastoma SH‐SY5Y cell line by Bennett et al. (1999). 6‐His‐tagged a‐synuclein constructs were transiently transfected and pulse–chase experiments revealed that the half‐life of the mutant form was 50% longer than that of the wild type. Degradation was inhibited by lactacystin‐b‐ lactone indicating that degradation was mediated by the proteasome, but there was no evidence for any polyubiquitinated forms. It was suggested that the greater propensity of mutant a‐synuclein to aggregate may be due to its slower clearance by the proteasome (Bennett et al., 1999). However, Ancolio et al. (2000) did not find proteasome‐mediated degradation of either a‐synuclein or the A53T mutant in HEK293 cells or in neuronal cells after transient transfection of these cells with constructs containing either a‐synuclein or the A53T mutant. Neither lactacystin nor the proteasome inhibitor PSI affected the levels of a‐synuclein. They also prepared stable transfectants in TSM1 neurons. Again the proteasome inhibitors did not modify immunoreactive a‐synuclein. They concluded that either the concentration of lactacystin‐b‐lactone used in the Bennett study was too high (40 mM) or that susceptibility to proteasome degradation is a cell‐specific phenomenon. In agreement with Bennett et al., they found no evidence of ubiquitinated a‐synuclein. Although a‐synuclein is a proteasome substrate in vitro, it is apparently degraded only very slowly in vivo. Its metabolic stability was explored by stably transfecting HK293 cells and PC12 cells with a‐synuclein cDNA and with a FLAG‐tagged molecule (Okochi et al., 2000). Pulse–chase analysis showed a‐synuclein to be a very stable protein. In addition, the protein was found to be constitutively phosphorylated most likely by casein kinases 1 and 2. A major phosphorylation site identified as Ser 129 lies within the C‐terminal domain. Ser 87 was also phosphorylated, but less efficiently. a‐Synuclein extracted from brains of patients with dementia with Lewy bodies is extensively phosphorylated on Ser 129 (Fujiwara et al., 2002). Greater than 90% of insoluble a‐synuclein in the affected brains was phosphorylated whereas less than 4% was phosphorylated in normal brains. Fujiwara et al. also found that phosphorylation promotes aggregation


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leading to their proposal that phosphorylation plays a role in the pathogenesis of neurodegenerative diseases. Full‐length phosphorylated a‐synuclein is mono‐ and di‐ubiquitinated (Hasegawa et al., 2002). It is unclear if this has anything to do with the ubiquitin/proteasome pathway since at least a tetraubiquitin chain is required for proteasome recognition. There is good evidence that the major pathway for a‐synuclein degradation is not proteasomal but rather lysosomal. Cuervo et al. (2004) working with rat ventral midbrain cultures confirmed the long half‐life of a‐synuclein (16.8 h). The proteasome inhibitor epoxomicin had only a minor effect on the half‐ life, whereas NH4Cl, a lysosomal inhibitor, significantly increased the half‐life. Wild‐type a‐synuclein was internalized into lysosomes and degraded by a chaperone‐mediated pathway. By contrast, mutant a‐synucleins were poorly internalized and therefore inefficiently degraded. Moreover mutant a‐synucleins blocked the uptake and degradation of other substrates of this pathway. Cuervo et al. (2004) propose that a‐synuclein may contribute to cell stress by blocking the degradation of other long‐lived proteins.

3.1.2 a‐Synuclein and the Proteasome The mechanism of interaction of a‐synuclein and the proteasome is controversial, with contradictory results reported in the literature. Martin‐Clemente et al. (2004) reported that overexpression of either wild‐ type or mutant a‐synuclein in mice or PC12 cells did not significantly affect proteasome activity, subunit expression, or assembly and function of the proteasome complex. Snyder et al. (2003) reported that aggregated a‐synuclein strongly inhibited the 26S proteasome (IC50 ¼1 nM), but inhibition of the 20S proteasome was relatively weak (IC50 > 1 mM). The selective strong inhibition of the 26S proteasome suggested that a‐synuclein interacts with a component of the 19S cap complex. Additional experiments showed that both aggregated and monomeric a‐synuclein bound to the S60 (Rpt5) subunit of the base of the 19S cap complex. This subunit is believed to mediate binding and recognition of ubiquitin (see > Sect. 2.4.2). By contrast, Lindersson et al. (2004) failed to detect binding of aggregated a‐synuclein filaments to the S60 subunit or to other 19S subunits but found binding to 20S proteasome subunits as well as holo 20S. Moreover the filaments selectively, noncompetitively, and potently inhibited the chymotrypsin‐like activity of the 20S proteasome (IC50 ¼1 nM). Lindersson et al. (2004) speculate that the discrepancies of the two studies may be due to different proteasome preparations or to different states of aggregation of a‐synuclein. Nonetheless, inhibition of the proteasome (either the 20S or 26S forms) by aggregated a‐synuclein could contribute to its neuropathology. The cytotoxicity of mutant a‐synuclein and its relationship to the proteasome was explored by Tanaka et al. (2001b). They established a PC12 cell model in which either a‐synuclein or the A30P mutant could be induced and found that the three catalytic activities of the proteasome measured in whole‐cell extracts were modestly decreased in cells expressing mutant a‐synuclein. These cells were also more susceptible to lactacystin‐induced cell death. Cells expressing mutant huntingtin did not show enhanced toxicity to lactacystin. In the mutant a‐synuclein‐expressing cells, there was also an increased sensitivity to mitochondria‐dependent apoptosis. These studies provide a link between proteasome impairment and mitochondrial dysfunction and suggest that this may have a causal role in Parkinson’s disease. There is evidence that the toxicity of a‐synuclein is related to lipid metabolism. The biological basis of a‐synuclein pathology was explored in a yeast cell model (Outeiro and Lindquist, 2003). A construct of a‐synuclein fused to green fluorescent protein under the control of a galactose‐inducible promoter was prepared and integrated into the yeast genome. The protein associated with the plasma membrane. Increasing its expression by integrating a second copy in the genome led to appearance of a‐synuclein in large cytoplasmic inclusions. Since some but not all aggregates had ubiquitin immunoreactivity, it was concluded that ubiquitination was neither necessary for aggregation nor prevented it. Notably, accumulation of a‐synuclein impaired the ability of the proteasome to degrade an unstable green fluorescent protein derivative. a‐Synuclein also inhibited phospholipase D and promoted lipid droplet accumulation. Inhibition of phospholipase D was found to be a biologically relevant and highly conserved property of a‐synuclein. A relationship of a‐synuclein to lipid metabolism was also reported by Willingham et al. (2003). They established a genomic screen in yeast to identify genes that could enhance the toxicity

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of a‐synuclein or of a mutant fragment of the huntingtin protein. When wild‐type or mutant A53T a‐synucleins were overexpressed in yeast, cytoplasmic inclusion bodies formed. Approximately 2% of the genes enhanced the toxicity of wild‐type a‐synuclein. Of the genes that have a known functional role, the highest percentage was associated with lipid metabolism. The primary source of toxicity of a‐synuclein may therefore not be related to the ubiquitin‐proteasome pathway. The genes regulating a‐synuclein toxicity and the toxicity due to the huntingtin fragment did not overlap, suggesting different pathogenic mechanisms for Parkinson’s disease and Huntington’s disease. The nature of a‐synuclein in Lewy bodies fractionated from human brain tissue was characterized by Tofaris et al. (2003). Urea extracts from brains of patients with Parkinson’s disease and from patients with diffuse Lewy body disease contained a‐synuclein immunoreactivity, whereas an extract from control brains did not. A specific species of a‐synuclein of 22–24 kDa was a substrate for mono‐, di‐, and tri‐ ubiquitination. Since the degradation signal for proteolysis by the ubiquitin‐proteasome system requires at least a tetraubiquitin chain, the ubiquitinated forms are unlikely to reflect targeting for degradation. The activity of the proteasome was reduced in some but not all of the affected regions. It was suggested that accumulation of this modified and partially phosphorylated 22–24 kDa form of a‐synuclein may overwhelm the proteolytic system, leading to aberrant ubiquitination and inclusion formation.

3.1.3 a‐Synuclein‐Interacting Protein: Synphilin‐1 A yeast two‐hybrid screen was used to identify proteins interacting with a‐synuclein (Engelender, 1999). A novel protein was identified and termed synphilin‐1. Although the physiological function of synphilin‐1 is not known, it contains several protein–protein interaction domains. It was proposed to act as an adaptor protein by anchoring a‐synuclein to intracellular proteins involved in vesicle transport and in the function of the cytoskeleton. Cotransfection of synphilin‐1 with constructs encoding parts of a‐synuclein led to the formation of Lewy body‐like cytoplasmic inclusions. It was suggested that the interaction between these two proteins could play a role in the formation of inclusion bodies in Parkinson’s disease. Synphilin‐1 is predominantly a neuronal protein and is enriched in presynaptic terminals. As discussed in > Sect. 3.3, there is evidence for ubiquitination of synphilin‐1 by parkin. More recently this ubiquitination was found to be nonclassical i.e., the ligation was to lysine 63 and not to lysine 48 (Lim et al., 2005). Ubiquitination of synphilin‐1 appears to be unrelated to degradation by the proteasome but may be related to Lewy body formation.

3.2 Parkin Mutations in a gene termed parkin appear to be responsible for ARJP (Kitada et al., 1998). This form of Parkinsonism, although displaying clinical features similar to the idiopathic form, differs in its absence of Lewy bodies and in its early onset, although there is evidence that mutations in the parkin gene may be of significance in some patients with late‐onset disease (Klein et al., 2000). The gene encodes a 465‐amino‐acid protein with an interesting structure. Parkin contains an ubiquitin‐like domain at its amino terminus and two ring finger motifs in its C‐terminal portion. The ring finger domains flank a cysteine‐rich domain termed ‘‘in‐between RING’’ or IBR. Proteins of the family RING1‐IBR‐RING2 recruit E2’s and serve as a novel type of E3. Evidence that parkin functions as an E3 was provided by Shimura et al. (2000) who demonstrated that parkin specifically interacted with the E2 UbcH7 via its ring finger domain and reacted more weakly with UbcH8. Parkin also binds ubiquitinated proteins before their degradation by the proteasome. An in vitro ubiquitination assay system provided direct evidence for the E3 activity of parkin. The molecular mechanisms whereby a parkin mutation causes ARJP are unknown. Tanaka et al. (2001a) proposed that either a yet to be determined proteasome substrate is not degraded leading to cell death or that increased levels of this proteasome substrate cause the cells to become more sensitive to environmental factors. Parkin appears to function as an E3, as part of a larger complex (Staropoli et al., 2003). It associates with the F‐box/WD repeat protein hSel‐10 and with cullin, a component of the SCF E3 ligase, to form a complex.


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The other SCF components Skp‐1 and Rbx1 do not associate indicating that the parkin E3 complex is unique. Cyclin E is a target substrate of this complex and increased cyclin E levels have been correlated with apoptosis. Embryonic primary cultures of midbrain DA neurons transfected with parkin siRNA were sensitized to kainate excitotoxicity. The overexpression of parkin protected these neurons from apoptosis and inhibited the accumulation of cyclin E. When cells are treated with lactacystin, parkin significantly accumulates in the perinuclear region and is colocalized with the centrosomal marker g tubulin (Zhao et al., 2003). The recruitment of parkin is dependent on intact microtubule networks and is disrupted by colchicine. Lactacystin also induces the accumulation of the parkin substrate CDcrel‐1 (see below) and ubiquitinated proteins in the centrosome. The significance of protein accumulation into a perinuclear structure termed the aggresome is more fully discussed in > Sect. 4.1. Zhao et al. (2003) suggest that parkin may ubiquitinate unfolded proteins accumulated in this region and speculate that parkin may also be necessary for the formation of Lewy bodies. This would explain why patients with ARJP‐harboring mutant parkin do not have Lewy bodies. It is of interest that parkin‐deficient mice do not display any clear Parkinsonism phenotype (Goldberg et al., 2003; Itier et al., 2003). There is no reduction in the number of dopaminergic neurons in the substantia nigra and the levels of several putative parkin substrates (see below) are not increased. However, the parkin‐deficient mice did display some mild deficits in dopaminergic function and some behavioral changes, suggesting some role for parkin in dopaminergic regulation (> Figure 13-2).

. Figure 13-2 Role of parkin in autosomal recessive juvenile Parkinsonism (ARJP). Parkin, an E3 ubiquitin–protein ligase normally targets its substrates for degradation by the 26S proteasome. Mutant parkin found in ARJP cannot function as an E3. It is proposed that the failure of a Parkin substrate(s) to be degraded is responsible for nigral cell degeneration (Tanaka et al., 2001)

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3.2.1 Search for the Parkin Substrate Investigators have sought to identify the substrate of parkin that could account for the pathological changes in ARJP. Several candidates have been proposed. One of these is the synaptic vesicle‐associated protein CDCrel‐1. Yeast two‐hybrid screening with parkin as bait identified CDCrel‐1 as an interacting protein (Zhang et al., 2000), and interaction was confirmed by coimmunoprecipitation experiments. Parkin also promoted the ubiquitination of CDCrel‐1 and enhanced its degradation. These investigators confirmed that parkin reacted with both UbcH7 and UbcH8, but point out that only UbcH8 is abundantly expressed in the CNS. They further found that parkin binds to UbcH8 via its R2 ring finger domain. Mutations within this domain disrupt the binding and impair the degradation of CDCrel‐1. Synaptogamin XI is another parkin‐interacting protein identified by the yeast two‐hybrid system (Huynh et al., 2003). Synaptogamin XI is a member of a family of calcium‐binding proteins and may play a regulatory role in signal transduction. As a result of this interaction, synaptogamin XI is ubiquitinated and degraded. Both parkin and synaptogamin XI colocalize in dense core vesicles. In contrast to parkin, synaptogamin XI was also found in Lewy bodies. The presence of parkin and synaptogamin XI in dense core vesicles led Hyunh et al. (2003) to speculate that the two proteins may play a role in calcium‐ dependent exocytosis and that loss of function of parkin may result in altered dopamine release. A two‐hybrid screen of a human brain cDNA library identified a homolog of endothelin receptor type B as a parkin‐interacting protein (Imai et al., 2001). This protein was renamed parkin‐associated endothelin receptor‐like receptor or Pael‐R. Parkin ubiquitinates Pael‐R and accelerates its degradation. When overexpressed, Pael‐R becomes unfolded, insoluble, and ubiquitinated and causes cell death. Parkin suppresses both the accumulation of Pael‐R and cell death. Imai et al. (2001) reported that the brains of patients with ARJP contained elevated levels of Pael‐R compared to controls and suggested that the accumulation of Pael‐R is causative in ARJP. There also is evidence that the ubiquitination of Pael‐R by parkin is facilitated by CHIP (Imai et al., 2002). Unfolded proteins are either refolded by chaperones or targeted to the proteasome for degradation. CHIP contains a U‐box motif, has E3 ligase activity, and ubiquitinates unfolded proteins when they are first captured by chaperones. When flag‐tagged parkin was immunoprecipitated and immunoprecipitates were analyzed by 2D gel electrophoresis and mass spectrometry, Hsp70 and CHIP were identified as binding partners. It was found that CHIP acting as an E4, potentiated the ubiquitination of Pael‐R by parkin, and facilitated its subsequent degradation by the proteasome. CHIP also promoted the dissociation of Hsp70 from parkin and Pael‐R. A fourth parkin‐interacting protein identified by the yeast two‐hybrid system is p38, a component of the aminoacyl‐tRNA synthase complex (Corti et al., 2003). In cells overexpressing parkin, p38 was ubiquitinated and its degradation was significantly enhanced. Loss of parkin function could result in an accumulation of nonubiquitinated p38. p38 is present in Lewy bodies of patients with Parkinson’s disease. These results provide a link between Parkinson’s disease and a protein involved as a structural component of a complex necessary for protein biosynthesis. Ren et al. (2003) reported that parkin binds to microtubules, that both a‐ and b‐tubulin can be coimmunoprecipitated with parkin, and that parkin is localized along microtubules. Overexpression of parkin increased the amount of polyubiquitinated a‐tubulin and accelerated its degradation. Furthermore parkin mutants linked to ARJP did not ubiquitinate tubulin. It is speculated that in the presence of mutant parkin, misfolded tubulin can accumulate and be toxic to cells.

3.3 a‐Synuclein and Parkin Since a‐synuclein is predominantly degraded by the proteasome in an ubiquitin‐independent manner and since parkin functions as an E3, the consequence of parkin‐mediated ubiquitination of a‐synuclein is unclear. Shimura et al. (2001) reported that ubiquitination does occur and that a complex of parkin, UbcH7 and a new isoform of a‐synuclein exists in the human brain. Immunohistochemistry revealed colocalization of parkin and a‐synuclein in Lewy bodies. The new isoform of a‐synuclein migrates at 22 kDa rather than at the expected 16 kDa (also see Tofaris et al., 2003; > Sect. 3.1.2). The 22 kDa form is due to O‐glycosidation


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since incubation with O‐glycosidase and sialidase shifts its migration to 16 kDa. Shimura et al. (2001) further reported that wild‐type parkin ubiquitinates the 22 kDa form of a‐synuclein in an in vitro system and that mutant parkin does not. They propose that loss of parkin results in an accumulation of nonubiquitinated 22 kDa a‐synuclein in ARJP. They further suggest that parkin could also play a role in idiopathic Parkinson’s disease. This proposal is based on the observations that wild‐type parkin mediates the ubiquitination of a‐synuclein as well as other protein substrates and that these proteins accumulate in Lewy bodies when the proteasome degradation system is impaired. By contrast, Chung et al. (2001) failed to confirm ubiquitination or interaction of the 22 kDa form of a‐synuclein with parkin. They instead found that synphilin‐1 was coimmunoprecipitated with parkin and that the immunoprecipitated synphilin‐1 was ubiquitinated. When synphilin‐1 and a‐synuclein were cotransfected, cytosolic inclusions were formed, and these inclusions were only rarely ubiquitinated. When parkin was coexpressed with these two proteins, the inclusions were ubiquitinated. They propose that it is the ability of synphilin‐1 to interact both with a‐synuclein and parkin that provides a link between these molecules and the pathogenesis of Parkinson’s disease. Petrucelli et al. (2002) suggest that vulnerability of cells to proteasome inhibition provides a link between parkin and a‐synuclein and a possible explanation of the selective sensitivity of a subgroup of dopaminergic neurons in Parkinson’s disease. M17 neuroblastoma cells transfected with mutant A53T a‐synuclein were much more sensitive to toxicity to proteasome inhibitors than cells transfected with wild‐ type a‐synuclein. Mutant a‐synuclein enhanced the toxicity of cells to proteasome inhibitors by decreasing proteasome function and toxicity could be partially antagonized by parkin. Proteasome activity in living cells monitored by the disappearance of a GFP‐fusion protein was inhibited by lactacystin, and the effect of lactacystin was enhanced by mutant a‐synuclein. Mutant a‐synuclein was also more toxic to tyrosine hydroxylase‐positive cells of primary midbrain cultures. This observation is of interest in view of the selective vulnerability of these cells in Parkinson’s disease. Caution needs to be exercised when conclusions are drawn concerning the impairment of the ubiquitin‐ proteasome system when a CMV‐driven expression system is used (Biasini et al. (2004)). The role of the ubiquitin‐proteasome system in the degradation of parkin and a‐synuclein under endogenous conditions was studied in PC12 cells transfected with parkin or a‐synuclein under the control of a CMV promoter. Exposure of the cells to either MG132 or lactacystin produced a time‐dependent increase in the levels of parkin and of a‐synuclein. The accumulation of these proteins appeared to be a consequence of increased synthesis rather than decreased degradation since Northern blot analysis showed a dramatic increase in the mRNA of both parkin and a‐synuclein following exposure to the proteasome inhibitors. Moreover protein synthesis inhibitors blocked the increase in protein levels. When the experiment was repeated in nontransfected cells, the proteasome inhibitors had no effect on endogenous levels of either parkin or on parkin mRNA and endogenous a‐synuclein could not be detected. Similarly, proteasome inhibitors did not increase endogenous parkin levels in primary mesencephalic cultures. Proteasome inhibitors may therefore increase CMV‐driven transcription.

3.4 Studies on the Proteasome in Parkinson’s Disease Several studies have directly addressed the question of the role of the proteasome in Parkinson’s disease by measuring proteasome activities in autopsy samples. A decrease of 33–42% in the catalytic activities of the proteasome in the substantia nigra of patients with Parkinson’s disease compared to control specimens was reported by McNaught and Jenner (2001). Although these studies were controlled for specificity by determining that the activity attributed to the proteasome was blocked by lactacystin, they must still be viewed with caution (Rogers and Dean, 2003). The specificity of lactacystin is questionable (see > Sect. 2.1.4). The substrate used to measure trypsin‐like activity is nonspecific and is cleaved by trypsin‐ like enzymes in the brain (Shibatani and Ward, 1995). Vigouroux et al. (2003) have partially purified a nonproteasomal peptidase in brain capable of degrading the most commonly used chymotrypsin‐like proteasome substrate, succinyl‐Leu‐Leu‐Val‐Tyr‐amc. We have confirmed their results and have identified this enzyme as thimet oligopeptidase (EC 3.4.24.15). Thimet oligopeptidase cleaves this substrate at internal

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sites, and the amc reporter group is released from the products by abundant brain aminopeptidases present in crude brain extracts. Several studies have sought to relate proteasome function to the selective loss of dopaminergic terminals in parkinsonism. When enriched fetal rat ventral mesencephalic neuronal cultures were exposed to lactacystin, a decrease in the number of dopaminergic terminals was observed, as was inhibition of dopamine uptake. (McNaught et al., 2002). There was also an increase in the staining of a‐synuclein in perikarya and processes and formation of proteinaceous cytoplasmic inclusions. Dopaminergic neurons were found to be more susceptible to lactacystin than GABAergic neurons. When proteasome inhibitors (lactacystin and epoxomicin) were infused into the striatum of rats, a selective neurotoxicity in dopaminergic neurons occurred (Fornai et al., 2003). The lesion side contained intracellular inclusions that stained for parkin, a‐synuclein, ubiquitin, and E1. Similar inclusion staining for the same components were found in PC12 cells treated with lactacystin. Agents that decreased dopamine protected against lactacystin‐ induced inclusions whereas agents that increased dopamine had an opposite effect. Fornai et al. (2003) proposed that dopamine may play a role in the lactacystin‐induced neurotoxicity perhaps by forming oxidation adducts. A dysfunction of the ubiquitin‐proteasome system may prevent the proper clearance of toxic intermediates and account for the selective vulnerability of dopaminergic neurons in Parkinson’s disease. Since the identification of the selective dopaminergic neurotoxicity of MPTPþ (Burns et al., 1983), there has been speculation that environmental toxins may underlie idiopathic Parkinson’s disease. McNaught et al. (2004) propose that environmental exposure to proteasome inhibitors may play a causative role. When the proteasome inhibitor PSI was administered to rats, the animals developed a gradually progressive motor dysfunction that resembled features of parkinsonism. There was a 53% loss of tyrosine hydroxylase‐positive neurons in the substantia nigra, a loss of tyrosine hydroxylase‐immunoreactive neurons in the striatum, and a depletion of dopamine and its metabolites. Inclusion bodies formed that stained positive for a‐synuclein, synphilin‐1, ubiquitin, tubulin, and parkin. These investigators propose that proteasome inhibition may be a good animal model for Parkinson’s disease. Their studies also uncover a possible long‐term toxicological problem in the use of proteasome inhibitors for cancer treatment (Adams, 2003). It is notable that a 12‐week exposure of SH‐SY5Y cells to low amounts (100 nM) of the nonselective proteasome inhibitor MG115 led to an increase in the levels of free ubiquitin, oxidized proteins, and aggregated proteins (Ding et al., 2003). However, proteasome inhibitors are neuroprotective in a 6‐hydroxydopamine model of Parkinson’s disease perhaps by facilitating inclusion formation and sequestering toxic components (Inden et al., 2005). The possibility that the Lewy body may be a specialized form of aggresome, present in dopaminergic neurons for protection against abnormal proteins, has been proposed (McNaught et al., 2001; Olanow et al., 2004). Thus Lewy bodies contain g‐tubulin, components of the ubiquitin‐proteasome pathway, heat‐shock proteins, and a‐synuclein. The Lewy body is proposed to initially play a protective role when first formed, but eventually becomes nonfunctional. The absence of Lewy bodies in ARJP may contribute to the early and severe cell death in this disorder (see > Sect. 4.2 for a discussion of aggresomes).

4

Protein Aggregation and the Proteasome

The presence of unfolded proteins within the cell can have deleterious effects on cell function. Unfolded proteins present exposed hydrophobic surfaces that are buried in the native conformer. Exposure of these hydrophobic chains to the aqueous environment can facilitate interactions with other unfolded proteins thereby leading to the formation of aggregates. Aggregated proteins that result from an expansion of polyglutamine residues may be responsible for the cytotoxicity of Huntington’s disease and the other polyglutamine expansion diseases (Perutz and Windle, 2001). To eliminate unfolded proteins, mechanisms have evolved to either refold the proteins by the assistance of chaperones or to target them for degradation by the proteasome. The presence of intracellular inclusions containing aggregated proteins is a common feature of many neurodegenerative diseases including Parkinson’s disease, the polyglutamine expansion


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diseases, and Alzheimer’s disease. Such inclusions contain immunoreactive ubiquitin and components of the ubiquitin‐proteasome system. Since unfolded proteins are normally cleared by the ubiquitin‐proteasome system, it is reasonable to assume that either inhibition of the proteasome by the aggregated proteins or an underlying defect in the ubiquitin‐proteasome system may be responsible for impaired degradation of the aggregates and formation of inclusions. However, whether the ubiquitin‐proteasome system is impaired in neurodegenerative diseases is the subject of considerable debate. Similarly, whether inclusions contribute to the neuropathology or are cytoprotective is also a matter of contention.

4.1 The Aggresome Wojcik et al. (1996) sought to determine whether treatment of HeLa cells with a proteasome inhibitor would lead to the accumulation of ubiquitin‐protein conjugates. HeLa cells were exposed for 24 h to 5 mM of the proteasome inhibitor PSI (Figueiredo‐Pereira, 1994). Heavily stained perinuclear aggregates generally located in a nuclear indentation around the centrosome were visible usually with one aggregate in each cell. The aggregates also stained with ubiquitin antiserum and with a monoclonal antibody directed against a proteasome subunit. Aggregate formation was blocked by cycloheximide. When the cells were treated with nocodazole, aggregate formation was not blocked but the aggregates were dispersed from the perinuclear region suggesting that they were connected to a microtubule scaffold. Western blotting of whole‐cell extracts showed an accumulation of ubiquitin‐containing complexes of high‐molecular weight. Wojcik et al. (1996) proposed that the aggregates represent ‘‘proteolysis centers’’ i.e., that the bulk of cellular proteolysis is spatially organized and concentrated in the perinuclear region. Johnston et al. (1998) studied the effect of proteasome inhibition on the disposition of the cystic fibrosis transmembrane conductance regulator (CFTR), an integral membrane protein. A deletion of phenylalanine 508 from CFTR (DF508CFTR) is the most common cystic fibrosis‐causing allele and the mutation impairs the ability of this protein to properly fold. When HEK cells expressing DF508CFTR were exposed to proteasome inhibitors, they observed large increases in aggregates of this protein. Similarly aggregates were formed when the protein was overexpressed. The protein was then tagged with GFP to examine its cellular disposition. Almost all of the GFP fluorescence was found in a single large perinuclear structure. This structure, which also stained with an antiubiquitin antibody, was termed ‘‘the aggresome.’’ There was a strong correlation with the staining of the protein and that of the centrosome marker g‐tubulin. Treatment with nocodazole abrogated accumulation of the protein at the aggresome, indicating that an intact microtubule network is essential for aggresome formation. Similar results were found for the integral membrane protein presenillin‐1. The aggresome was defined as an inclusion body that is formed by retrograde transport of aggregated proteins on microtubules (Kopito, 2000). It was proposed that aggresome formation is a general cellular response to an overwhelming of the capacity of the proteasome either due to excess substrate or reduced proteolytic activity. Retrograde transport on microtubules may serve to clear the cell of potentially toxic aggregates and the delivery of aggregated proteins into aggresomes may be a means of increasing the efficiency of their capture for autophagy (Kopito, 2000). To investigate the functional significance of aggresomes, a reporter consisting of a C‐terminal destabilizing sequence (Gilon et al., 1998) fused to GFP was prepared (Bence et al., 2001). This fusion protein was much less stable than GFP, was ubiquitinated, and its degradation blocked by a proteasome inhibitor. Fluorescence of this construct in cells was used as a reporter of activity of the ubiquitin proteasome system. Fluorescence was measured in cells transiently expressing the aggregation prone protein DF508CFTR. When this protein was expressed, aggresomes formed and fluorescence of the GFP reporter increased. Protein aggregation led to accumulation of intracellular ubiquitin conjugates and cell cycle arrest. It was concluded that formation of protein aggregates could lead to a decreased activity of the ubiquitin proteasome system and consequent neurodegeneration. Donaldson et al. (2003) proposed that ubiquitin‐rich aggregates serve as a site for the recruitment and sequestration of ubiquitin‐binding proteins and that depletion of ubiquitin‐binding proteins may contribute to neurodegeneration. Additional evidence for a role of the centrosome in protein degradation was provided by the studies of Wigley et al. (1999) in HEK293 cells. They employed immunocytochemistry to determine the location of

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proteolysis in the cell . Although various components of the ubiquitin proteasome pathway and chaperones were widely distributed, 20S proteasome, 19S regulator complex (PA700), 11S REG (PA28), ubiquitin, and the centrosomal marker g‐tubulin all colocalized in a unique perinuclear site. Centrosomes were then purified by density gradient centrifugation and subjected to Western blot analysis. All of the above components copurified with the centrosome. When cells were treated with lactacystin, there was a significant increase in the size of the centrosome and the size of the centrosome could also be increased by the expression of misfolded proteins. The aggregated proteins were apparently recruited to the centrosome for degradation (> Figure 13-3).

. Figure 13-3 Aggresome formation. Misfolded proteins that are not degraded by the proteasome either because the capacity of the proteasome is overwhelmed or because its activity is impaired may spontaneously aggregate. The aggregates are delivered via microtubules to the perinuclear aggresome for sequestration and degradation


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4.2 Polyglutamine Diseases The polyglutamine diseases including Huntington’s disease are a group of disorders in which neurodegeneration results from a protein containing an expanded polyglutamine tract. Huntington’s disease is a progressive autosomal dominant neurodegenerative disorder with both motor and cognitive components. Patients with Huntington’s disease exhibit chorea, spasticity, and progressive dementia. In 1993, the Huntington’s disease collaborative research group reported on a mutation in the first exon of a gene containing an unstable CAG repeat and encoding a 350‐kDa protein of unknown function (Huntington’s Disease Collaborative Research Group, 1993). In normal patients, this gene product termed huntingtin contains a poly‐Q repeat of 8–37 residues, whereas in patients with Huntington’s disease the expansion is 38–182 residues. Other neurodegenerative disorders containing a poly‐Q expansion include spinobulbar muscular atrophy also known as Kennedy’s disease, dentatorubral‐pallidoluysian atrophy, and the spinocerebellar ataxias SCA1, SCA2, SCA3 (also known as Machado‐Joseph disease), SCA6, SCA7, and SCA17. Spinobulbar muscular atrophy is X‐linked whereas the others are dominantly inherited. The function of several of the proteins causing the poly‐Q disorders is known. Spinobulbar muscular atrophy is due to a poly‐Q expansion of the androgen receptor (La Spada et al., 1991). SCA6 is due to a poly‐Q expansion in the a1A voltage‐dependent calcium channel (Zhuchenko et al., 1997), and SCA17 is due to a poly‐Q expansion in the TATA‐binding protein, a general transcription initiation factor (Nakamura et al., 2001). The neuropathological profile presented is dependent on the protein containing the expanded poly‐Q tract. The proteins involved in the poly‐Q diseases show no homology apart from the poly‐Q tracts, and neurodegeneration can be induced by introducing an expanded poly‐Q tract into a nontoxic protein. Thus, when a 146Q tract was inserted into the gene for hypoxanthine phosphoribosyltransferase, mice harboring this mutation developed a delayed onset and progressive neurological phenotype (Ordway et al., 1997). Studies on transgenic flies expressing poly‐Q tracts alone demonstrated that these tracts are intrinsically neurotoxic (Marsh et al., 2000). In general, once the poly‐Q tract exceeds 36–40 residues, there is an earlier appearance of symptoms and progressive neurodegeneration that is proportional to the length of the repeat. Perutz at al. (1994) have presented evidence that the poly‐Q repeats act as polar zippers joining molecules together. They have shown by molecular modeling that hydrogen bonding links together two antiparallel poly‐Q strands. To support this hypothesis, they prepared Asp2Gln15Lys2 as a model poly‐Q peptide and demonstrated that this peptide formed aggregates of tightly linked b‐sheets. They further proposed that neurodegeneration is due to an aggregation and precipitation of these proteins in neurons.

4.2.1 Colocalization of Inclusions and Components of the Ubiquitin‐Proteasome System Mice transgenic for the Huntington’s disease mutation were generated by Davies et al. (1997). These mice harbor CAG repeats of 115–156 units and develop a progressive and complex neurological phenotype. The brains of these mice contained NII. These densely stained circular inclusions are found within neuronal but not glial cells, and contain immunoreactive huntingtin and ubiquitin. The inclusions are found in almost all striatal neurons. Ultrastructural analysis of the striatum showed indentations of the nuclear membrane and an increase in clustering and number of nuclear pores. These investigators suggested that the NII are the primary site for cellular dysfunction. Studies of patients with poly‐Q diseases have documented that the NII contain elements of the ubiquitin‐proteasome system. For example, immunoreactive ubiquitin and immunoreactive 20S proteasome subunits colocalize with mutant ataxin‐1 aggregates in brain sections from patients with SCA1 (Cummings et al., 1998). In brainstem sections of patients with SCA3 (Machado‐Joseph disease), NII were immunoreactive for ubiquitin, 20S proteasome, 19S regulator complex, and 11S REG (PA28) (Chai et al., 1999). Ubiquitin colocalized with NII from the brain of a patient with SCA7 (Holmberg et al., 1998) and colocalized with huntingtin in NII and dystrophic neurites in brain sections from Huntington’s disease patients (DiFiglia et al., 1997). Motor neurons of spinobulbar muscular atrophy patients contained

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androgen receptor‐immunoreactive ubiquitinated nuclear inclusions (Li et al., 1998), and molecular chaperones were also found in these inclusions (Cummings et al., 1998). The brains and spinal cords of seven patients with dentatorubral‐pallidoluysian atrophy contained ubiquitin‐immunoreactive NII (Hayashi et al., 1998). NII in a patient with SCA17 also stained with ubiquitin antibodies and with antibodies against the TATA‐binding protein carrying a 48 CAG repeat (Nakamura et al., 2001). However, NII were not seen in SCA2 brain tissues (Hyunh et al., 1999). This may be due to the shorter length of the poly‐Q tract in mutant ataxin‐2 compared to the other poly‐Q disease proteins. Also three brains of patients with SCA6 did not have immunoreactive ubiquitin inclusions (Ishikawa et al., 1999). Again the poly‐Q tract in these patients was relatively short (22 residues).

4.2.2 Interaction of Proteasome and Poly‐Q‐Containing proteins Evidence for interaction of the proteasome and poly‐Q‐containing proteins in general was provided by Nollen et al. (2004), who sought to identify genes which when suppressed could lead to premature appearance of protein aggregates. They developed a transgenic Caenorhabditis elegans model that expressed poly‐Q expansions as yellow fluorescent protein‐fusion proteins. In this screen, RNAi bacteria were fed to C. Elegans in liquid culture and animals visually scored for foci formation. Among the 186 genes identified were genes encoding the 20S subunit a‐4, a subunit of the 19S regulatory complex, E1, and a member of the ubiquitin family. Ataxin‐3 Hofmann and Falquet (2001) searched for proteins containing potential ubiquitin‐binding

sequences by selecting a ‘‘ubiquitin‐interacting region’’ present in subunit 5a (Rpn10) of the 19S regulatory complex. They found that the ubiquitin‐interacting motif (UIM) occurred in a wide variety of proteins. One of the identified proteins was ataxin‐3 which contains a tandem UIM upstream of the poly‐Q sequence and an additional UIM downstream of the poly‐Q sequence in a splice variant. Immunoprecipitation and pull‐down experiments established that ataxin‐3 is indeed an ubiquitin‐ binding protein (Burnett et al., 2003; Chai et al., 2004). Both normal and poly‐Q expanded forms of ataxin‐3 coimmunoprecipitate ubiquitinated proteins (Chai et al., 2004). Ataxin‐3 binds ubiquitin tetramers but not ubiquitin monomers or dimers, and its poly‐Q tract is not necessary for binding. Ataxin‐3 is also able to bind proteasome subunits (Doss‐Pepe et al., 2003). Ataxin‐3 may be recruited to polyubiquitin‐containing inclusions because of its ubiquitin‐binding properties. Of interest is the finding that ataxin‐3 can also function as a ubiquitin protease (Burnett et al., 2003). There is sequence homology around cysteine 14 and histidine 119 of human ataxin‐3 and the active site residues of ubiquitin proteases. Ataxin‐3 deubiquitinates ubiquitin‐125I‐lysozyme and mutation of Cys14 to Ala abolishes this activity. Ataxin‐3 stabilizes some short‐lived proteins (Burnett et al., 2003; Doss‐Pepe et al., 2003), presumably by its deubiquitinating activity. The functional significance and relationship to disease pathology of the ubiquitin‐binding properties and the deubiquitinating activity of ataxin‐3 remain to be determined. Ataxin‐3 also interacts with the polyubiquitin chain elongation factor E4B and with the multiubiquitin‐chain‐binding protein VCP (valosin‐containing protein) perhaps as a ternary complex (Matsumoto et al., 2004). Pulse–chase experiments in the presence and absence of a proteasome inhibitor established that ataxin‐3 is a proteasome substrate and that the presence of a 79Q tract markedly slows degradation. The yeast two‐hybrid system was used to identify proteins interacting with huntingtin. The ubiquitin‐conjugating enzyme E2–25K was identified and the two proteins were also shown to interact in an in vitro binding assay. It is of interest that interaction was not dependent on the length of the poly‐Q tract. The huntingtin protein itself was found to be ubiquitinated suggesting that it is a substrate of the ubuiquitin‐proteasome system. The high expression of E2–25K in brain suggests that E2 participates in huntingtin ubiquitination in vivo (Kalchman, 1996).

Huntingtin


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4.3 The Role of the Proteasome in Protein Aggregation and Inclusion Formation Since components of the ubiquitin‐proteasome system are found in NII, the functional significance of this association is of considerable interest. Cummings et al. (1999) hypothesized that the poly‐Q expansion in ataxin‐1 caused the protein to misfold, and thereby be targeted for degradation by the proteasome. They proposed that inclusions form as the capacity of the proteasome is exceeded. This hypothesis was supported by studies in which HeLa cells were transfected with an 82Q ataxin‐1 fusion protein. When the transfected cells were treated with lactacystin, there was a great facilitation of ataxin‐1 aggregation and an increase in the size of the nuclear aggregates. Significantly, the distribution of the cellular 20S proteasome was also altered in that the proteasome was redistributed to the aggregates. A significant amount of the mutant ataxin protein was found in the detergent‐insoluble fraction of the cell and the detergent‐insoluble material was ubiquitinated. Proteasome inhibition increased the levels of the detergent‐insoluble form but not the soluble form. A cell‐free system was established and it was shown that mutant ataxin‐1 was degraded by the proteasome less efficiently than the normal form. To investigate the role of a compromised ubiquitin‐ proteasome system on SCA1 transgenic mice, mice deficient in E6‐AP, an E3 (see > Sect 2.6.4), were crossed with the SCA1 transgenic mice. Although the double mutant mice developed fewer nuclear aggregates, examination of brain sections showed more severe pathology. It was concluded that the formation of nuclear inclusions is not necessary for neurodegeneration. These studies indicate that ataxin‐1 is a substrate for the proteasome and that the mutant poly‐Q‐expanded form is degraded more slowly. The true E3 responsible for targeting ataxin‐1 for degradation remains to be identified and the investigators were unable to demonstrate an E6‐AP‐dependent ubiquitination of ataxin‐1 in vitro. However, it remains possible that the pathology was exacerbated in the double mutant independent of the failure of ataxin‐1 to be degraded but rather as a consequence of the failure of another target of E6‐AP to be ubiquitinated and degraded. A role for the proteasome in aggregate formation is also supported by the studies of Chai et al. (1999). Transfection of HeLa or COS-7 cells with a truncated Q49-ataxin-3 construct led to a general diffuse staining pattern with Sect. 4.3 indicate that degradation of poly‐Q‐containing proteins by the proteasome is inversely proportional to the length of the poly‐Q expansion. Venkatraman et al. (2004) show that purified 26S proteasomes cannot degrade within poly‐Q sequences in vitro. Thus, a Q20 peptide containing an RRGRR flanking sequence for solubility was cleaved by the 26S proteasome only within the flanking sequence. These investigators also studied open‐channel forms of the 20S proteasome and found that even the open‐channel forms could not degrade within a poly‐Q stretch. Their studies suggest that poly‐Q‐containing proteins can be degraded by the proteasome outside the poly‐Q tract, thereby generating an aggregation prone poly‐Q fragment that may play a role in the neuropathology. In their model, proteasomes may promote neurotoxicity by forming products that are more toxic than their substrates. They further suggest that long poly‐Q tracts may have difficulty diffusing out of the proteasome and may therefore act as proteasome inhibitors. Inhibition of the proteasome would lead to a buildup of abnormal proteins (but at the same time limit the formation of the toxic poly‐Q peptides). In cells expressing a 150Q huntingtin fragment, there is a reduction in proteasome activity and in the degradation of the proteasome substrate p53 (Jana et al., 2001). Since poly‐Q chains do not normally accumulate, they should be substrates of another yet to be identified proteinase(s). Venkatraman et al. (2004) suggest that such an enzyme(s) may play an important role in clearance of these potentially toxic products. In experiments using fluorescence live‐cell imaging, Holmberg et al. (2004) showed that overexpression of a construct of N‐terminal huntingtin containing a Q65 repeat caused a redistribution of proteasome to aggregates. They also reported that poly‐Q‐containing proteins were partially degraded by the proteasome. These proteins may also, in part, be trapped within the proteasome leading to impaired proteasome function. Impaired proteasome function is also observed when ataxin‐1 (82Q) is transfected into cells suggesting that this impairment contributes to the pathology (Park et al., 2005). However, some studies do not confirm proteasome inhibition under conditions in which a poly‐Q‐ expanded protein is expressed (Bennett et al., 2005), and there are also reports of efficient degradation of poly‐Q‐containing proteins by the proteasome. Apparently targeting is essential for the proteasome to degrade poly‐Q‐containing proteins. Verhoef et al. (2002) linked poly‐Q repeats containing 16, 65, or


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112 residues to the C terminus of an ubiquitin‐R‐GFP proteasome substrate. Ubiquitin is cleaved by a C‐terminal hydrolase exposing R at the N terminus and thereby creating an N‐end rule substrate. Cells were transfected with the construct or an N‐terminal Met‐containing control protein. The ubiquitin‐R‐GFP protein was degraded by the proteasome and insertion of the poly‐Q repeat did not significantly slow degradation. This demonstrates that proteins containing a poly‐Q repeat can be degraded by the proteasome provided a degradation signal is present. Proteins recruited to nuclear inclusions were resistant to degradation. Thus, poly‐Q proteins are stable once they aggregate and formation of aggregates can be slowed by introduction of a degradation signal into the protein. Michalik and Broeckhoven (2004) also showed that the proteasome can efficiently degrade model substrates containing either 103‐ or 25Q provided that the substrate contains an N‐end rule degradation signal. Proteasome activities in brains of mice harboring a 94Q expansion of huntingtin under the control of a CamKIIa promoter in a tetracycline‐regulated manner were determined by Diaz‐Hernandez et al. (2003). They found no evidence for inhibition, but an increase in the chymotrypsin‐like and trypsin‐like activities of the proteasome compared to controls in the cortex and striatum of these mice. There was no significant increase in proteasome content as judged by Western blotting. Ding et al. (2002) using a different model also showed that a poly‐Q expansion did not impair proteasome activity. Both groups reported an upregulation of immunoproteasome subunits. Diaz‐Hernandez et al. (2003) reported that Western blotting showed a significant increase in the immunoproteasome subunits LMP2 and LMP7. They also showed intense staining of LMP2 in degenerating neurons in the brains of HD94 mice. The physiological significance of the upregulation of the immunoproteasome remains to be determined. A decrease in proteasome enzymatic activity in fibroblasts and in several brain regions of patients with Huntington’s disease was reported by Seo et al. (2004). They feel that the increases described in the mouse transgenic model may represent a compensatory mechanism that may occur in early‐onset or severe forms of the disease. Problems associated with direct measurement of proteasome activities in tissues due to the nonspecific nature of the substrates was discussed in > Sect. 3.4.

5

Conclusion

It is clear from the above discussion that the relationship between the ubiquitin‐proteasome system and the poly‐Q expansion neurodegenerative diseases remains unresolved. Many elegant models have been developed to probe this possible association, but it has not been determined whether an impairment of the ubiquitin‐proteasome system underlies these diseases or even whether reduced proteasome activity contributes to the pathology. It is not clear if protein aggregates directly decrease proteasome activity since evidence for and against this effect is reported in the literature. What can account for these contradictory results? This may in part be due to the common experimental approach of transfection of supraphysiological amounts of substances in an attempt to derive information about physiological processes. In some cases, accumulation of protein is interpreted as a decrease in degradation but may be due to an increase in transcription. The presence of ubiquitin and proteasome components in aggregate‐containing inclusions could be due to an underlying defect in the ubiquitin‐proteasome system or may mean that aggregated proteins are poor substrates of a normal ubiquitin‐proteasome system. Delivery of aggregated proteins into aggresome‐like inclusions appears to be a protective mechanism of the cell since there is abundant evidence that dispersed microaggregated species are cytotoxic. It is also possible that the focus on the ubiquitin‐proteasome system is misplaced. The suggestion that yet to be identified proteolytic enzymes are responsible for clearing poly‐ Q tracts is of considerable interest (Venkatraman et al., 2004) and alterations in a nonproteasome proteinase may contribute to the neuropathology of the poly‐Q diseases. Perhaps the best evidence for a causal relationship of the ubiquitin‐proteasome system to neurodegeneration is in ARJP where parkin was identified as an E3. Although mutations in parkin are linked to ARJP, there is no clear parkinsonian phenotype in parkin‐null mice, nor is there any link of putative parkin substrates to Parkinson’s disease. a‐Synuclein is degraded by the proteasome in an ubiquitin‐independent manner and its turnover in the cell is relatively slow. The major pathway for degradation of this protein

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appears to be lysosomal rather than proteasomal. Although a viable ubiquitin‐proteasome system is essential for normal cell function, the role of this important protein degradation pathway in Parkinson’s disease remains to be determined.

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does not correlate with the formation of intranuclear inclusions. Cell 95: 55-66. Scheffner M, Huibregtse JM, Howley PM. 1994. Identification of a human ubiquitin‐conjugating enzyme that mediates the E6‐AP‐dependent ubiquitination of p53. Proc Natl Acad Sci USA 91: 8797-8801. Schmidt M, Haas W, Crosas B, Santamaria PG, Gygi SP, et al. 2005. The HEAT repeat protein Blm10 regulates the yeast proteasome by capping the core particle. Nat Struct Mol Biol 12: 294-303. Seemuller E, Lupas A, Stock D, Lowe J, Huber R, et al. 1995. Proteasome from Thermoplasma acidophilum: A threonine protease. Science 268: 579-582. Seo H, Sonntag KC, Isacson O. 2004. Generalized brain and skin proteasome inhibition in Huntington’s disease. Ann Neurol 56: 319-328. Sheaff RJ, Singer JD, Swanger J, Smitherman M, Roberts JM, et al. 2000. Proteasomal turnover of p21Cip1 does not require p21Cip1 ubiquitination. Mol Cell 5: 403-410. Shibatani T, Ward WF. 1995. Sodium dodecyl sulfate (SDS) activation of the 20S proteasome in rat liver. Arch Biochem Biophys 321: 160-166. Shimura H, Hattori N, Kubo S, Mizuno Y, Asakawa S, et al. 2000. Familial Parkinson disease gene product, parkin, is a ubiquitin–protein ligase. Nat Genet 25: 302-305. Shimura H, Schlossmacher MG, Hattori N, Frosch MP, Trockenbacher A, et al. 2001. Ubiquitination of a new form of a‐synuclein by parkin from human brain: Implications for Parkinson’s disease. Science 293: 263-269. Snyder H, Mensah K, Theisler C, Lee J, Matouschek A, et al. 2003. Aggregated and monomeric a‐synuclein bind to the S60 proteasomal protein and inhibit proteasomal function. J Biol Chem 278: 11753-11759. Sorimachi H, Kawasaki H, Tsukahara T, Ishiura S, Emori Y, et al. 1991. Sequence comparison among subunits of multicatalytic proteinase. Biomed Biochim Acta 50: 459-464. Spillantini MG, Schmidt ML, Lee VM, Trojanowski JQ, Jakes R, et al. 1997. a‐Synuclein in Lewy bodies. Nature 388: 839-840. Staropoli JF, McDermott C, Martinat C, Schulman B, Demireva E, et al. 2003. Parkin is a component of an SCF‐like ubiquitin ligase complex and protects postmitotic neurons from kainate excitotoxicity. Neuron 37: 735-749. Sudakin V, Ganoth D, Dahan A, Heller H, Hershko J, et al. 1995. The cyclosome, a large complex containing cyclin‐ selective ubiquitin ligase activity, targets cyclins for destruction at the end of mitosis. Mol Biol Cell 6: 185-197. Tanahashi N, Murakami Y, Minami Y, Shimbara N, Hendil KB, et al. 2000. Hybrid proteasomes. Induction by interferon‐g and contribution to ATP‐dependent proteolysis. J Biol Chem 275: 14336-14345.

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14

Serine Proteases

J. To´th . P. Medveczky . L. Szila´gyi . L. Gra´f

1

The Structure and Function of Serine Proteases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410

2 2.1 2.2 2.3 2.4 2.5

Plasma Serine Proteases in the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 413 Activity and Specificity of Hemostatic Serine Proteases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 413 Chromosomal Localization of Hemostatic Serine Proteases and Regulation of their Expression 414 Characterization of the Protease‐Activated Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 414 Localization of Hemostatic Serine Proteases and PARs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 415 The Role of Plasma Serine Proteases in the Physiology and Pathology of the CNS . . . . . . . . . . . . . 416

3 3.1 3.2 3.3 3.4 3.5 3.6

Kallikreins in the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418 Nomenclature, Genomic Localization, and Structure of Kallikrein Genes . . . . . . . . . . . . . . . . . . . . . . . . 418 Regulation of Expression and Alternative Splicing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 419 Structure and Specificity of Kallikrein Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425 Biological Roles of Kallikreins and Their Association with Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425 Potential Roles of Kallikreins in the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 Kallikreins in Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427

4 4.1 4.2 4.3 4.4

Human Trypsin 4—an Unconventional Serine Protease in the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . 428 Chromosomal Localization and Transcription of Human Trypsin Genes . . . . . . . . . . . . . . . . . . . . . . . . 428 Regional Distribution of Human Trypsin 4 mRNA and Protein in the Brain . . . . . . . . . . . . . . . . . . . . 429 Structural and Biochemical Characterization of Human Trypsin 4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 431 On the Potential Role of Human Trypsin 4 in Physiological or Pathological Processes in the CNS . . 433

5

Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 435

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Serine proteases

Abstract: Serine proteases are an ancient and large group of proteolytic enzymes that utilize a uniquely activated serine residue to catalyze peptide bond hydrolysis. Members of this protease family are highly diversified and are involved in numerous physiological and pathological processes. Their enzymatic activities are tightly regulated through transcription, translation, zymogen activation, autolysis, and interaction with natural inhibitors. In this chapter, the authors will begin with a brief description of the structure and function of serine proteases. Next, several members of this family of enzymes, including thrombin, plasmin, plasminogen activators, kallikreins, and human trypsin 4 are presented. These sections will discuss the genomic localization, gene structure, and regulation of expression of these proteases followed by the description of their enzymatic specificity and biological functions focusing on processes in the central nervous system (CNS). List of Abbreviations: 50 ‐RACE, rapid amplification of 50 ‐cDNA ends; AA, amino acid; AD, Alzheimer’s disease; AP1, activating protein 1; AP2, activating protein 2; APP, amyloid precursor protein; BPTI, bovine pancreatic trypsin inhibitor; cAMP, 30 ,50 ‐cyclic adenosine monophosphate; CNS, central nervous system; CSF, cerebrospinal fluid; EAE, experimental autoimmune encephalomyelitis; ELISA, enzyme‐linked immunosorbent assay; EPO, erythropoietin; FTD, frontotemporal dementia; GDN, glia‐derived nexin; GFAP, glial fibrillar acidic protein; GFP, green fluorescent protein; GnRH, gonadotropin‐releasing hormone; GPCR, G‐protein coupled receptor; HBE, human bronchial epithelial cell line; HEK, human embryonic kidney cell line; HIF, hypoxia‐inducible factor; hK, human kallikrein; hPSTI, human pancreatic secretory inhibitor; HRE, hypoxic response element; IGFBP‐3, insulin‐like growth factor‐binding protein‐3; kcat, catalytic constant; KLK, kallikrein gene; Km, Michaelis constant; LTP, long‐term potentiation; mAb, monoclonal antibody; MALDI‐MS, matrix‐assisted laser desorption ionization mass spectrometry; MBP, myelin basic protein; MOG, oligodendrocyte glycoprotein; MS, multiple sclerosis; MSA, multiple system atrophy; MSP, myelencephalon specific protease; MUGB, 4‐methylumbelliferyl 4‐guanidinobenzoate; NF1, nuclear factor 1; NMDA, N‐methyl‐D‐aspartate; PAI‐1, plasminogen activator inhibitor 1; PAR, protease‐activated receptor; PCR, polymerase chain reaction; PD, Parkinson’s disease; PN1, protease nexin I; PRSS, serine protease gene; RT‐PCR, reverse transcription polymerase chain reaction; SP1, signal protein 1; STI, soybean trypsin inhibitor; TCR, T‐cell receptor; TGF‐b, transforming growth factor b; TM, thrombomodulin; tPA, tissue‐type plasminogen activator; uPA, urokinase‐type plasminogen activator; uPAR, urokinase‐type plasminogen activator receptor

1

The Structure and Function of Serine Proteases

Proteases (also termed proteinases, peptidases, or proteolytic enzymes) are a huge and important group of enzymes that hydrolyze peptide bonds (an excellent review of the field is the Handbook of Proteolytic Enzymes, 2004). Proteases are grouped on the basis of primary and tertiary structures into families and clans, which are further grouped based on catalytic mechanisms. This classification was introduced by Rawlings and Barrett (1995). A family of proteases is a group of enzymes in which every member shows a significant relationship in the amino acid (AA) sequence of the catalytic domain to at least one other member of the family. In other words, any two members of a family have evolved from a common ancestor and are homologous. The families are named with a letter denoting the catalytic type followed by an arbitrary number. A group of families form a clan in which the members have evolved from a common ancestor but have diverged to such an extent that their relationship cannot be revealed by comparing their primary structures (Rawlings and Barrett, 1993; Barrett and Rawlings, 1995). The name of the clan is composed of the letter for the catalytic type followed by an arbitrary second letter. Proteases reviewed here (thrombin, plasmin, plasminogen activators, trypsin, and kallikreins) are endopeptidase serine proteases belonging to clan SA, family S1. Endopeptidases act internally in polypeptide chains and they can be further divided on the basis of their catalytic mechanism. Five catalytic types of endopeptidases can now be recognized, in which serine, threonine, cysteine, aspartic, or metallo groups play primary roles in catalysis. The serine, threonine, and cysteine proteases are catalytically very different from the aspartic and metallopeptidases in that the nucleophile of the catalytic site is part of an amino acid, whereas it is an activated water molecule in the other groups.

Serine proteases

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In the S1 family of proteases the catalytic mechanism depends upon the hydroxyl group of a serine residue acting as the nucleophile that attacks the peptide or amide bond, and thus they are termed serine proteases. The residues indispensable for their enzymatic activity, His57, Asp102, and Ser195, (chymotrypsinogen numbering) are structurally conserved and are referred to collectively as the catalytic triad (Huber, 1978). These residues are located at the active site of the enzyme. As the first step of the catalytic cycle, the enzyme binds the substrate forming the noncovalent Michaelis complex. The hydroxyl oxygen of the catalytic serine makes a nucleophilic attack on the electron‐deficient carbonyl carbon of the scissile bond and a covalent bond is formed. The attacked carbon atom becomes tetrahedrally coordinated; thus, this state is called the first tetrahedral intermediate. In the next step, this species breaks down to yield the C‐terminal (first) product and the acyl‐enzyme. The acylation is followed by deacylation, a mechanistically similar action, where the acyl–ester bond is attacked by a water molecule that has previously been activated by His57 (Bender and Ke´zdy, 1965; Polga´r and Bender, 1969). During the hydrolysis of the scissile bond tetrahedral intermediates are developed which are stabilized via hydrogen bonding interactions, where the amido groups of residues Gly193 and Ser195 (forming the oxyanion hole) act as H‐donors and the developing oxyanion intermediate is the acceptor (Henderson, 1970; Robertus et al., 1972). Most of these proteases enter the secretory pathway and have an N‐terminal signal peptide, and all of them are synthesized in zymogen forms as precursors with an N‐terminal extension namely the propeptide, to be clipped off to generate the active enzyme. In trypsin, the most intensively studied member of family S1, the cleavage of the propeptide leads to the disruption of a His40–Asp194 hydrogen bond, followed by the rotation of Asp194 allowing the interaction with the newly formed N terminus at Ile16. This conformational change is triggered by the formation of a salt bridge between the a-amino group of Ile16 and the side chain carboxylate group of ASP 194 and leads to the completion of oxyanion hole and subtrate-binding pocket formation (Fehlhammer et al., 1977; Bode et al., 1978). The structure has been solved for chymotrypsinogen and trypsinogen, the precursors of chymotrypsin and trypsin, respectively (Kraut, 1971; Bode et al., 1984; Wang et al., 1985), and the comparison between the structures of the proenzyme and the active enzyme is striking. The part of the molecule rearranged during this conformational change is called the ‘‘activation domain.’’ The new conformations are stabilized mainly by hydrophobic interactions of the Ile16 side chain. (Hedstrom et al., 1996). It was proposed on theoretical grounds, that the ‘‘activation domain’’ of serine proteases might govern both zymogen activation and catalysis (Gra´f, 1995). Unique conformational differences were revealed between the human plasminogen and plasmin structures that are not seen in other zymogen–enzyme pairs of the trypsin family (Wang et al., 2000). The proenzyme of human kallikrein (hK) 6 displays a fold that exhibits chimeric features between those of trypsinogen and other family members: it possesses a completely closed specificity pocket and a unique conformation of the regions involved in structural rearrangements upon activation. This points to a novel activation mechanism, which could be extrapolated to other hKs (Gomis‐Ruth et al., 2002). The length of the propeptide varies greatly. The smallest one belongs to human cathepsin G that contains two residues, and the larger ones contain hundreds of amino acids. The larger propeptide region, found for example in the blood coagulation factors and complement components, commonly remain attached by disulfide bonds to the protease domain after proteolytic activation, and the extension is referred to as the ‘‘heavy chain’’ of the heterodimeric active enzyme. The N‐terminal propeptide extension may be mosaic in nature, containing domains that have had separate evolutionary origins from the protease domains. The highly conserved three‐dimensional structure of the catalytic residues provides an extremely efficient framework for peptide bond cleavage. Tertiary structures have been solved for a number of proteases of family S1. > Figure 14-1 shows the structure of human trypsin 4. The chymotrypsin fold shows a two‐domain structure with an active site cleft between the domains. Each domain consists of a b‐barrel and the two barrels are arranged at right angles to each other. It has been suggested that the two‐ domain structure is the result of a gene duplication and fusion event, and that the ancestral gene possessed only one b‐barrel domain (Lesk and Fordham, 1996). The residues indispensable for the catalytic activity, His57, Asp102 and Ser195, are structurally conserved and located in the active site cleft between the two domains. Ser195 acts as a nucleophile during the peptide (or amide) bond cleavage, His57 serves as a general base (Kossiakoff and Spencer, 1981), while Asp102 stabilizes the correct tautomer of His57 and compensates the developing positive charge during the catalytic reaction.

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Serine proteases

. Figure 14-1 Three‐dimensional structure of human trypsin 4 in complex with benzamidine based on the Protein Data Bank file 1h4w visualized using software PyMOL (DeLano, 2002). Residues His57, Asp102, and Ser195 comprising the catalytic triad are displayed along with Arg193 located in the C terminus of Loop1 and the benzamidine molecule in the substrate‐binding pocket

The substrate peptide forms an antiparallel b‐sheet with the substrate‐binding site. The catalytic site is flanked by specific subsites that are able to interact with the appropriate residues of the substrate peptide. The nomenclature for these subsites was introduced by Schechter and Berger (Schechter and Berger, 1967). These sites are numbered from the active site (S1) as S2, Sn toward the N terminus, and S20 , Sn0 toward the C terminus (Sn‐S2‐S1‐S10 ‐S20 ‐Sn0 ). The respective residues on the substrate peptide are numbered Pn‐P2‐ P1‐P10 ‐P20 ‐Pn0 . The substrate specificity for trypsin‐type proteases is primarily determined by residue 189 (chymotrypsinogen numbering) at the bottom of the S1 binding pocket, which is an Asp for trypsin. The residues at position 216 and 226 (glycines for trypsin) are also important for the specific interaction. A disulfide bridge between Cys191 and Cys220 and a loop formed by residues 214–220 also determine the structure of the substrate‐binding pocket. Furthermore, it was shown that loop1 (residues 185–193) and loop2 (residues 217–224) also affect the specificity. Trypsin‐type proteases possess six disulfide bonds in conserved positions (Cys 15–145, 33–49, 117–218, 124–191, 156–170 and 181–205). The autolysis loop is comprised of residues 143–151. The proteolytic activity of proteases is under strict regulation in several ways including transcriptional regulation, zymogen activation, autolysis, and interaction with endogenous inhibitors. Additional structural domains might also occur in the N‐terminal extensions in the S1 family of endopeptidases, e.g., apple domains, kringle domains, finger domains, epidermal growth factor‐like domains, cysteine‐rich domains, etc. Several of these possess the characteristic patterns of disulfide bonds. The acquisition of these additional domains is thought to have been the result of exon shuffling during evolution (Patthy, 1990). This was made possible by the presence of a phase 1 intron–exon junction near the 50 ‐end of the protease domain in the ancestral gene, which is retained in many present day genes for proteases of family S1.

Serine proteases

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Serine proteases possess widely diverse biological functions in animals such as the digestion of foodstuff, cascade‐like activation of other proteases, cleavage and activation of specific receptors and growth factors, coagulation and fibrinolysis, processing of peptide hormones, tissue remodeling, and degradation of extracellular matrix proteins.

2

Plasma Serine Proteases in the CNS

This part of the chapter is focused on a class of serine proteases, namely the hemostatic enzymes, which include thrombin, plasmin, urokinase‐type plasminogen activator (uPA), and tissue‐type plasminogen activator (tPA). Much is known about hemostatic proteases within the coagulation system, but we are just beginning to understand the important role that these proteases play in neuronal cell development and pathology. There are many pathologies of the central nervous system (CNS) due to injury or disease in which blood may enter the neural parenchyma and potentially affect neural and glial cells. Besides their important role in signaling, thrombogenesis, and fibrinolysis, hemostatic proteases are also involved in tumor cell invasion, inflammation, oocyte maturation, cell migration, and in many other processes in CNS biology (reviewed by Dano et al., 1985; Turgeon and Houenou, 1997; Gingrich and Traynelis, 2000; Rohatgi et al., 2004; Suo et al., 2004; Sheehan and Tsirka, 2005).

2.1 Activity and Specificity of Hemostatic Serine Proteases Thrombin, one of the most prominent members of the serine protease family, is a 36‐kDa protein comprised of two chains, A and B, linked by a disulfide bond. The A‐chain has 36 amino acid residues whereas the B‐chain, which contains the serine protease domain, has 259 residues (Mann, 1994). Thrombin is derived from its proenzyme or zymogen form (prothrombin or coagulation factor II) through cleavage by factor Xa in the presence of cofactor Va, Ca2þ, and a phospholipid surface (Krishnaswamy et al., 1993). Prothrombin and thrombin were among the first subjects of many protein chemical enzymatic studies in the early 1980s. The nucleotide sequence for prothrombin was also determined in the 1980s (Degen et al., 1983). The elucidation of the tertiary structure of thrombin (Bode et al., 1989) led to a number of groups focusing on thrombin as a target for drug development (Tapparelli et al., 1993). The P1 residue of substrates and inhibitors of thrombin is almost always arginine, and the specificity is generally trypsin‐like, but much more restricted. The restriction of specificity results from (1) partial occlusion of the active site, restricting access of macromolecular substrates, (2) a greater selectivity for P3, P2, P20 and P30 residues, and (3) the use of extended interaction areas, termed exosites, distant from the active site (Bode et al., 1989, 1992). The most abundant specific natural substrate of thrombin is fibrinogen. Thrombin triggers clot formation by releasing two oligopeptides (fibrinopeptides A and B) from the N‐termini of the a and b chains of fibrinogen (Higgins et al., 1983). The activity of thrombin is also important in stabilizing the fibrin clot; it activates factor XIII that in turn cross‐links the fibrin monomers (Lorand et al., 1993). Thrombin amplifies its own production by activating essential cofactors in the coagulation cascade (factors V and VIII) (Davie et al., 1991) and by catalyzing platelet activation, which provides the surface necessary for the assembly of the protease–cofactor complexes of the cascade. The plasminogen activators uPA and tPA act further along the coagulation pathway and their primary role is to activate plasminogen to form plasmin. Plasmin is yet another serine protease that operates in conjunction with fibrin to form a clot. uPA was first identified in urine and subsequently in human seminal fluid, plasma, and certain cancers (Dano et al., 1985). This enzyme has limited substrate specificity, cleaving the sequence Cys‐Pro‐Gly‐Arg560/Val561‐Val‐Gly‐Gly‐Cys that constitutes a small disulfide‐bridged loop in plasminogen. Despite the highly restricted substrate specificity of uPA, it has been demonstrated to hydrolyze a number of other proteins in vitro including fibronectin, fibrinogen, diphtheria toxin, and possibly uPA itself. These are poor substrates for uPA and are probably of no biological significance. Interestingly uPA’s own cellular receptor has been shown to be a relatively good substrate (Hoyer‐Hansen et al., 1992).

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Serine proteases

tPA converts plasminogen to the two‐chain serine protease plasmin by hydrolysis of a single Arg561/ Val562 peptide bond. The activation of plasminogen by tPA, both in the presence and absence of fibrin, follows Michaelis–Menten kinetics. Although different kinetic parameters have been reported, most authors agree that the activation rate of plasminogen increases several 100‐fold in the presence of fibrin. Thus, tPA is a relatively poor enzyme in the absence of fibrin, but the presence of fibrin strikingly enhances its activity (Lijnen et al., 1994).

2.2 Chromosomal Localization of Hemostatic Serine Proteases and Regulation of their Expression The gene for the prothrombin zymogen is localized on chromosome 11 in the human genome, and includes 14 exons interrupted by 13 introns (Royle et al., 1987). The human uPA gene is 6.4 kb comprising 11 exons and 10 introns, located on chromosome 10 (Degen et al., 1987). Comparison of the murine uPA gene with the porcine and human uPA genes reveals a higher sequence similarity even in the introns and flanking sequences. The transcription initiation site is flanked by common RNA polymerase II promoter elements, including a TATA box and a potential transcription factor‐binding site (Degen et al., 1987). Transcription of the gene produces a single mRNA form of 2.5 kb. uPA gene expression is upregulated by a wide variety of cytokines, growth factors, and hormones in many cells in a cell type‐specific manner. On the other hand the human tPA gene, located on chromosome 8, consists of 14 exons. Its exon–intron organization strongly suggests exon shuffling, whereby the distinct structural domains are encoded by a single exon or by adjacent exons (Lijnen and Collen, 1991). The proximal promoter sequences in the human tPA gene containing typical TATA and CAAT boxes and potential recognition sequences for transcription factors (e.g., activating proteins AP1 and AP2, nuclear factor 1 (NF1), and signal protein 1 (SP1)) have been identified (Feng et al., 1990; Kooistra et al., 1991). Consensus sequences of a 30 ,50 ‐cyclic adenosine monophosphate (cAMP)‐ responsive element and of an AP2‐binding site have also been reported, which may have a cooperative effect on constitutive tPA gene expression (Medcalf et al., 1990). Allelic dimorphism has been observed in the human tPA gene as a result of an Alu insertion/deletion event which occurred early in evolution (Ludwig et al., 1992).

2.3 Characterization of the Protease‐Activated Receptors Thrombin is the ultimate serine protease in the coagulation cascade and is known as coagulation factor IIa. Thrombin’s effect on neuronal and nonneuronal cells has attracted increasing attention when the first thrombin receptor, now known as protease‐activated receptor 1 (PAR1) was sequenced and cloned. With subsequent identification of other PAR family members thrombin’s effect on neural cells in development and disease of the nervous system are being intensively studied. To date, cDNAs encoding four PAR receptors (PAR1–4) have been identified. Three of these genes (human PAR1, PAR2, PAR3) map to a single chromosome (5q13) in a gene cluster (Kahn et al., 1998), while human PAR4 maps to chromosome 19p12. Among the PARs, PAR2 represents a class of trypsin/tryptase receptors, whereas PAR1, PAR3, and PAR4 are most effectively activated by thrombin (> Figure 14-2). PAR1 can also be maximally activated by Factor Xa, trypsin, and plasmin, whereas Factor VIIa, tPA, cathepsin G, elastase, and chymotrypsin do not activate these receptors (Ishihara et al., 1997). Interestingly, plasmin can also cleave PAR1 at a linker domain that lies downstream of the SFLLRN activation sequence, thereby inactivating the receptor (Kuliopulos et al., 1999). PAR1, PAR2, and PAR4, but not PAR3, can be activated by a short peptide (SFLLRN for PAR1, SLIGKV for PAR2, TFRGAP for PAR3, and GYPGQV for PAR4) corresponding to the first few N‐terminal residues revealed by proteolysis. Because PAR‐selective peptides can be designed, these peptides provide an important tool for evaluating the involvement of specific PARs in various processes (Hollenberg et al., 1997). The classic G‐protein‐associated pathways that could be involved in transducing thrombin action include: activation of phospholipase C, the release of arachidonic acid, inhibition of adenylate cyclase, inositol phospholipid hydrolysis, and increase in

Serine proteases

14

. Figure 14-2 The four protease‐activated receptors (PARs) belong to the G‐protein‐coupled receptor family. PARs possess seven transmembrane helices and trigger signal transduction pathways upon being cleaved by serine proteases. The activation cleavage site in their extracellular domain is specific for the PAR isoforms. The tethered ligand domains become exposed upon cleavage and bind to the cleaved receptors, which are thus activated. PARs can also be activated by peptides composed of specific sequences. The sequences of the newly formed N‐termini are displayed along with sequences in parentheses representing peptides that activate PARs more specifically than the native tethered ligand. PAR3 does not trigger signal transduction, at least not in endothelial cells. This figure was originally published by B. W. Festoff (2004)

intracellular calcium (Siess et al., 1989; Vu et al., 1991). Quite early after the initial cloning of the first thrombin receptor (Vu et al., 1991), now termed PAR1, studies of thrombin signaling in neural cells and the localization of PARs in the CNS began.

2.4 Localization of Hemostatic Serine Proteases and PARs Prothrombin was long thought to be synthesized exclusively in the liver, but using polymerase chain reaction (PCR), Northern blot analysis, and in situ hybridization techniques, Dihanich et al. (1991) have shown that prothrombin is also expressed in the developing and adult rat brain. Levels of prothrombin mRNAs are low in the neonatal rat brain and increase significantly by adulthood. After birth, prothrombin mRNA and protein is expressed in the olfactory bulb, cortex, superior colliculus, inferior colliculus, corpus striatum, and thalamus (Dihanich et al., 1991). Expression of prothrombin mRNA has also been shown in human astroglial cultures (Deschepper et al., 1991). Even though it is still not clear whether prothrombin is converted into thrombin within the CNS, these observations suggested important roles for prothrombin/ thrombin in the brain. Citron and coworkers reported the expression of prothrombin in skeletal muscle and also the detection of high‐level expression of message and active thrombin after birth (Citron et al., 1997). They also found that prothrombin was expressed in the murine (rat, mouse) spinal cord (Citron et al., 2000), and that both the message and protein, as well as active thrombin, were present in the neonatal spinal cord up to postnatal day 30. The presence of tPA mRNA, protein, and its proteolytic activity are observed in neurons and glia cells across brain regions including neocortex, hippocampus, hypothalamus, amygdala, and cerebellum (Soreq and Miskin, 1981, 1983; Sappino et al., 1993; Ware et al., 1995; Tsirka et al., 1997; Salles and Strickland, 2002). In the hippocampus, tPA immunoreactivity was found exclusively in the mossy fiber pathway. In the amygdala, tPA immunoreactivity was confined to the central and medial amygdala and was almost completely absent in the basal amygdala (Pawlak et al., 2003). In contrast, the expression of uPA is extremely low in the postnatal brain (Sappino et al., 1993; Ware et al., 1995). There are a number of genes containing hypoxic response element (HRE) in their promoter regions (O’Rourke et al., 1997), and the expression of these genes are regulated by a hypoxia‐inducible factor (HIF)

415

416

14

Serine proteases

(Semenza et al., 2000; Semenza 2001). These include typical hypoxia‐associated genes such as erythropoietin (EPO), thrombomodulin (TM), protease nexin I (PN1), and plasminogen activator inhibitor I (PAI‐1). PAI‐1 and PNI are upregulated by thrombin (Rao et al., 1990; Ho et al., 1994; Festoff et al., 1997). Through this thrombin increases HIF level under normoxic conditions (Festoff et al., 2001; Page et al., 2002). The first thrombin receptor cloned, PAR1, is widely distributed in neurons and glia (Weinstein et al., 1995; Niclou et al., 1998). PAR1 mRNA is widespread in prenatal rat brain tissue, and becomes more pronounced and confined to particular cell types in adult animals. Using in situ hybridization, PAR1 mRNA was detected in distinct cell layers of the cortex, subiculum, hypothalamus, thalamus, pretectum, ventral mesencephalon, cerebellum, and olfactory bulb. In the hippocampus, PAR1 transcripts can be found in the granule cell layer of the dentate gyrus, with more diffuse hybridization in the pyramidal cell layer of CA1. Intense hybridization was observed in Purkinje cells of the cerebellum and some brainstem nuclei (Weinstein et al., 1995). PAR2 immunoreactivity has been reported in cultured hippocampal neurons (Smith‐Swintosky et al., 1997). In addition, PAR3 can be found at low levels in the mouse brain (Ishihara et al., 1997).

2.5 The Role of Plasma Serine Proteases in the Physiology and Pathology of the CNS Irrespective of whether hemostatic proteases enter the nervous system through the blood–brain barrier or are synthesized by cells within the brain, these proteases may contribute to the pathophysiology of the CNS. During neural development, these serine proteases contribute to cell migration, neurite outgrowth, and synapse elimination. In adults, they play a key role in the regulation of neuronal survival and structural plasticity associated with learning and memory processes (> Figure 14-3). Their mechanism of action probably involves degradation of extracellular matrix components, activation/inhibition of PARs, or activation/inhibition of other proteases. The presence of tPA appears to be necessary for long‐term potentiation (LTP) maintenance in CA1,3 hippocampal pathways (Huang et al., 1996; Baranes et al., 1998). Another important role for tPA in the maintenance of LTP is that tPA facilitates N‐methyl‐D‐aspartate (NMDA) receptor‐mediated signaling by cleaving one of the NMDA receptor subunits (Nicole et al., 2001). Thrombin also facilitates LTP and enhances NMDA receptor‐mediated responses in CA1 neurons (Gingrich et al., 2000). This effect of thrombin could be inhibited by a selective thrombin antagonist hirudin and mimicked by the peptide agonist SFLRN via activation of PAR1 (Gingrich et al., 2000). Migration of neurons is a critical phase of brain development. This process has been proposed to be tPA dependent. Plasmin is converted from plasminogen by plasminogen activators. Fibrinolytic activity of plasmin is relatively high at birth and declines during the third postnatal week (Soreq and Miskin, 1981). In neural development at the embryonic stage tPA and uPA may be involved in different ways in neuronal growth (Sumi et al., 1992). Nerve growth cones at the tips of growing neurites facilitate neurite outgrowth and migration of neurons through the surrounding tissue. Fibrin–agarose zymography clearly shows that tPA is the main plasminogen activator associated with growth cone structures in postnatal rat brains (Garcia‐Rocha et al., 1994). Other results indicate that tPA accelerates activity‐dependent axonal rearrangement in adult hippocampus (Wu et al., 2000). Structural changes in synaptic connections are in close relationship with the formation of long‐term memory and learning. A recent study shows that blocking tPA activity by PAI‐I or knocking out the tPA gene in rat dramatically impairs the runway task training. This suggests that the proteolytic activity of tPA may facilitate synaptic plasticity during motor learning (Seeds et al., 2003). The urokinase plasminogen activator system consists of uPA, its glycolipid‐anchored receptor (urokinase‐type plasminogen activator receptor (uPAR)), and its two serpin inhibitors plasminogen activator inhibitor‐1, ‐2 (PAI‐1, ‐2). The urokinase plasminogen activator system is involved in cell signaling, interactions with integrins, invasion angiogenesis, and cell motility. The uPAR plays a role in the migration of different cell types and evidence is accumulating that this migration is mediated through integrins (Blasi and Carmeliet, 2002; Resnati et al., 2002). This system has been correlated with malignancy and brain tumor progression. CNS tumors overexpress components of the urokinase plasminogen activator system

Serine proteases

14

. Figure 14-3 The role of the plasminogen activator–plasminogen–plasmin system (a) and thrombin (b) in the central nervous system (see text for details). This figure was originally published by Gingrich MB and Traynelis SF (2000)

and their expression level correlates with the tumor grade (Yamamoto et al., 1994a, b; Mohanam et al., 1999, 2002). The molecular processes that lead to cell death in the CNS following brain trauma and cerebrovascular insult are under intense investigation. Hemostatic serine proteases trigger plenty of effects in neuronal and glial cells that are frequently associated with brain damage. Glial proliferation is often associated with brain injury, and the proliferating glia form a barrier to regenerating axons. Picomolar concentrations of thrombin, as well as the PAR1‐agonist peptide, stimulate astrocyte proliferation and reverse astrocyte stellation in culture (Cavanaugh et al., 1990; Grabham and Cunningham, 1995; Koh et al., 2005). Several

417

418

14

Serine proteases

proteins appear to be rapidly phosphorylated on tyrosine residues following PAR1 activation in cultured astrocytes. Furthermore, the tyrosine kinase inhibitor, herbimycin A, and the kinase inhibitor, staurosporine, can block thrombin‐mediated cell proliferation (Grabham and Cunningham, 1995; Debeir et al., 1996). Thrombin has also been proposed to upregulate glial expression of TM in in vitro models of glial injury (Pindon et al., 2000). Thrombin appears to trigger edema, which can damage both white and gray matter (Lee et al., 1996; Xi et al., 1998). Thrombin‐induced edema is potentiated by tPA, a thrombolytic protease, which can compete for endogenous thrombin inhibitors such as protease nexin 1 (Figueroa et al., 1998). Evidences now suggest that PAR1 activation can induce cell death in hippocampal cultures and motoneurons in avian spinal cord; cell death could be blocked by caspase inhibitors (Donovan et al., 1997; Turgeon et al., 1998; Turgeon et al., 1999). Thrombin‐induced apoptosis was blocked by tyrosine kinase inhibitors and inhibitors of RhoA, a GTP‐binding protein known to be involved in apoptotic pathways. On the basis of thrombin’s ability to precipitate seizures (Lee et al., 1997), mice engineered to lack PN1 have an increased susceptibility to kainic acid‐induced seizures and showed other signs of hyperexcitability (Luthi et al., 1997). Surprisingly, mice engineered to oversynthesize PN1 also had an increased susceptibility to kainic acid‐induced seizures (Luthi et al., 1997). These data indicate that the balance between serine proteases and serpin inhibitors might control neuronal excitability. If the balance between serine proteases and serpins becomes compromised there may be a deleterious effect on CNS neurons and glia. High levels of thrombin and prothrombin have been localized to senile plaques in brains of patients with Alzheimer’s disease (AD), suggesting a role for this protease in the pathogenesis of the disease (Cavanaugh et al., 1990; Nelson and Siman, 1990; Arai et al., 2006).

3

Kallikreins in the CNS

3.1 Nomenclature, Genomic Localization, and Structure of Kallikrein Genes The term ‘‘kallikrein’’ comes from the Greek word ‘‘kallikreas’’ meaning pancreas. It was originally used for the description of an enzyme found in the pancreas that cleaves kininogen to yield kinin, a hypotensive peptide (Bhoola et al., 1992). The kallikrein enzymes are divided into two major groups based on their molecular weight, substrate specificity, type of kinin released, and gene structure as plasma and tissue kallikreins (Movat, 1979). Recently, the term ‘‘tissue kallikrein’’ is used to describe a group of enzymes having highly conserved structures both at the gene and protein levels. Thus, although these enzymes are all termed kallikreins, they do not necessarily possess kininogenase activity. Tissue kallikreins were recently reviewed by Yousef et al. (2005). The plasma kallikrein gene is localized on chromosome 4q35. In contrast, the human tissue kallikreins are localized on chromosome 19q13.4 (Gan et al., 2000). Tissue kallikreins are serine proteases belonging to a family of genes. Analysis of the 19q13.4 genomic region revealed 13 putative functional serine protease genes along with five truncated pseudogenes and a nonfunctional cyclophilin A gene. The uniform nomenclature of these serine proteases have recently been established as various names were used for the same protein according to its possible function or tissue localization. The genes are denoted as KLK (kallikrein gene) and the protein as hK (human kallikrein protein) (Diamandis et al., 2000a). Furthermore, the naming system for serine protease genes using the prefix PRSS (Protease, serine) also stands for kallikreins. Alternative names for hKs are summarized in > Table 14-1. Like other trypsinogen‐like serine proteases, all kallikrein genes in the 19q13.4 region are formed of five coding exons and some of them also have one or more extra 50 ‐untranslated exons. The sizes of the individual exons are very similar for the different kallikrein genes, which indicates a close evolutionary relationship. In contrast, the sizes of the introns are quite different. The most accepted theory is that the kallikrein gene family evolved via gene duplication and exon shuffling (Nelson et al., 1999; Yousef and Diamandis, 2000). The genomic organization and sequence of kallikrein genes suggest that they arose from multiple rounds of gene amplification events. The overall sequence conservation among kallikreins, including the signal peptide, activation peptide, and the protease domain, is around 30–40% at both the nucleotide and

Serine proteases

14

. Table 14-1 Alternative names for human kallikrein genes Name KLK1 KLK2 KLK3 KLK4 KLK5 KLK6 KLK7 KLK8 KLK9 KLK10 KLK11 KLK12 KLK13 KLK14 KLK15

Alternative names hPRK: Human pancreatic/renal kallikrein hGK‐1: Human glandular kallikrein 1 PSA: Prostate‐specific antigen Prostase, KLK‐L1: kallikrein‐like protein 1, PRSS17, ARM1, EMSP1: enamel matrix serine protease 1 KLK‐L2: kallikrein‐like protein 2, HSCTE: human stratum corneum tryptic enzyme Protease M, neurosin, zyme, PRSS9 HSCCE: Human stratum corneum chymotryptic enzyme, PRSS6 Neuropsin, ovasin, TADG‐14: tumor‐associated differentially expressed gene 14 KLK‐L3: Kallikrein‐like protein 3 NES1: Normal epithelial cell specific 1, PSSSL1 TLSP: Trypsin‐like serine protease, hippostasin, PRSS20 KLK‐L5: Kallikrein‐like protein 5 KLK‐L4: Kallikrein‐like protein 4 KLK‐L6: Kallikrein‐like protein 6 Prostinogen, HSRNASPH, ACO, ACO protease: Alzheimer brain cortex serine protease

amino acid levels. In contrast, KLK1, 2, and 3 (called the ‘‘classical’’ kallikreins) share about 80% sequence identity. Furthermore, the sequence of the promoter region, the introns, the 30 ‐ and 50 ‐noncoding regions, and the pseudogenes in their promoter regions also show significant homology. This indicates that these genes were generated via very recent gene duplications events (Gan et al., 2000). The degree of conservation is approximately 30–40% between kallikrein protease sequences and trypsinogens from human (Emi et al., 1986) or chicken (Wang et al., 1995). In contrast, the conservation of trypsinogen sequences between these species is around 70%, which is much higher than the conservation among kallikreins in humans. It suggests that kallikrein genes are under much less functional constraint or evolutionary pressure when compared with trypsinogen genes.

3.2 Regulation of Expression and Alternative Splicing The expression of several kallikreins is under steroid hormonal regulation, including KLK2, KLK3 (Shan et al., 1997), KLK4 (Nelson et al., 1999), and KLK6 (Yousef et al., 1999). In the case of KLK2 and KLK3, the cis‐acting hormone response elements have also been identified (Cleutjens et al., 1996). The expression profiles have been determined in various tissues for most of the kallikreins by means of Northern blotting and reverse transcription polymerase chain reaction (RT‐PCR) as summarized in several papers (Katz et al., 1998; Harvey et al., 2000; Clements et al., 2001; Yousef and Diamandis, 2001, 2002b; Bernett et al., 2002; Diamandis and Yousef, 2002). In general, kallikreins tend to show coexpression in given tissues indicating that they might form an enzymatic cascade. Here, we focus only on the expression of kallikrein genes in the CNS. The mRNAs of all kallikrein genes except for KLK2, 3, and 15 were reported to be detected in the CNS by Northern blotting or RT‐PCR. The expression levels of KLK6‐12 and KLK14 seem to be higher in the CNS relative to other kallikrein genes. The transcription pattern of kallikrein genes in different areas of the brain is summarized in > Table 14-2 Alternative splicing is frequent within human tissue kallikrein genes. This process plays a significant role in development, physiology, and disease. The first extensive review on this subject was published recently (Kurlender et al., 2005) in which a systematic analysis of all currently known kallikrein alternative transcripts is presented including alternative splicing patterns, along with identification of 82 different kallikrein gene transcript forms and 56 different protein forms for KLK1–15. In the kallikrein locus, the majority of alternative splicing events occur within the protein‐coding region and to a lesser

419

Cerebellar white matter

Brain stem

Basal ganglia

Amygdala

1

3

4

5 (R) Yousef and Diamandis (1999)

(R) Scarisbrick et al. (2001) (R) Scarisbrick et al. (1997) (NF) Petraki et al. (2001); J. To´th, M. Palkovits, L. Gra´f, and L. Szila´gyi (unpublished results)

6 (R) Yamashiro et al. (1997); Little et al. (1997)

7 (R) Yousef et al. (2000c) (R) Yoshida et al. (1998a) (R) Mitsui et al. (1999)

8 (R) Yoshida et al. (1998a)


10 (P) Yousef et al. (2003)

11

12 (R) Yousef et al. (2000a)

13

(R) Yousef et al. (2001b)

14 (R) Yousef et al. (2001b)

14

Fetal brain

KLK Brain

. Table 14-2 Expression pattern of kallikreins in the central nervous system

420 Serine proteases

Formatio reticularis

CSF

(P) Melegos et al. (1997)

(R) Shimizu‐ Okabe et al. (2001)

(P) Yousef et al. (2003)


(NF) Shimizu‐ Okabe et al. (2001)

Cerebral cortex


(R) Yousef and Diamandis (1999) (P) Yousef et al. (2003)

Cerebellum

(P) Diamandis et al. (2000b); Okui et al. (2001); Diamandis et al. (2004) (NF) Petraki et al. (2001); J. To´th, M. Palkovits, L. Gra´f, and L. Szila´gyi (unpublished results)

(P) Diamandis et al. (2004)


Yousef et al. (2000c)

Yousef et al. (1999)


(R)

(R)

(P) Yousef et al. (2003); Diamandis et al. (2004)


(NF) Diamandis et al. (2004)



(NF) Diamandis et al. (2004)



Yousef et al. (2001b)


Yousef and Diamandis (2000) (R) Shimizu‐ Okabe et al. (2001)

Mitsui et al. (1999) (R) Mitsui et al. (1999); Shimizu‐ Okabe et al. (2001) (NF) Diamandis et al. (2004)

(R)

(R)

(R)

(R)

Serine proteases

14 421

Hippo campus


1


3


4


5

(R) Scarisbrick et al. (2001); Scarisbrick et al. (1997); Shimizu‐ Okabe et al. (2001); Petraki et al. (2001); J. To´th, M. Palkovits, L. Gra´f, and L. Szila´gyi (unpublished results)

6 (R) Petraki et al. (2001); J. To´th, M. Palkovits, L. Gra´f, and L. Szila´gyi (unpublished results)


7

Yoshida et al. (1998a); Mitsui et al. (1999); Shimizu‐ Okabe et al. (2001)

(R) Mitsui et al. (1999)

8


9

(R) Shimizu‐ Okabe et al. (2001);

10

(R) Yoshida et al. (1998b); Mitsui et al. (2000); Shimizu‐ Okabe et al. (2001)

11

(NF) Shimizu‐ Okabe et al., 2001

12

(R) Shimizu‐ Okabe et al., 2001

13

(R) Shimizu‐ Okabe et al., 2001

14

14

Frontal lobe

KLK Frontal cortex

. Table 14-2 (continued)


Putamen

Nucleus pontinus

Nucleus caudatus

(R) Petraki et al. (2001); J. To´th, M. Palkovits, L. Gra´f, and L. Szila´gyi (unpublished results) (NF) Petraki et al. (2001); J. To´th, M. Palkovits, L. Gra´f, and L. Szila´gyi (unpublished results) (R) Petraki et al. (2001); J. To´th, M. Palkovits, L. Gra´f, and L. Szila´gyi (unpublished results)

Serine proteases

14 423

1

3

4

(R) ¼ mRNA, (P) ¼ protein, (NF) ¼ Not Found

Substantia nigra

KLK Spinal cord

(P) Yousef et al. (2003)


(R) Scarisbrick et al. (2001) Scarisbrick et al. (1997)

6 (R) Yousef et al. (1999); Scarisbrick et al. (2001); Scarisbrick et al. (1997) Yousef et al. (2000c)

7 (R)

8

Yousef and Diamandis (2000)

9 (R)

10

11

12

13


14 (R)

14

. Table 14-2 (continued)


Serine proteases

14

extent in the 50 ‐untranslated regions (UTRs). The most common alternative splicing event is exon skipping (35%) and the least common events are cryptic exons (3%) and internal exon deletion (3%). 76% of kallikrein splice variants predicted to encode truncated proteins are the result of frameshifts. Accumulating evidence suggests that alternative kallikrein forms could be involved in many pathological processes or could be applied as biomarkers.

3.3 Structure and Specificity of Kallikrein Enzymes The common structural features of kallikreins at the DNA, mRNA, and protein levels have been reviewed by Yousef and Diamandis (2003a, 2003b). All kallikrein proteins are predicted to be synthesized as a preproenzyme bearing a 17–20 amino acid hydrophobic signal peptide, followed by a 4–9 (or longer in the case of hK5) amino acid activation peptide, ensued by the protease domain comprised of 223–238 amino acid residues. The signal peptides indicate that these enzymes are secreted serine proteases. Positions of the residues comprising the catalytic triad (His, Asp, Ser) are conserved. The active site is located in the cleft between the two b‐barrel domains. The residue at the bottom of the substrate‐binding pocket is either aspartate (for hK1, 2, 4, 5, 6, 8, 10, 11, 12, 13, 14) indicating trypsin‐like specificity or another residue (glutamate for hK15, asparagine for hK7, and glycine for hK9, indicative of a chymotrypsin‐like specificity, or serine for hK3). Classical kallikreins (hK1, 2, 3) possess 10 while the rest of tissue kallikreins have 12 cysteine residues in conserved positions forming 5 or 6 disulfide bonds as in other serine proteases (Kauffman, 1965). hK1, 2, and 3 possess a characteristic loop called the ‘‘kallikrein loop,’’ which is an insertion of 11 residues in front of the catalytic aspartate residue. Alternatively, shorter insertions can be found at the same position in hK12 (two residues), hK8 (three residues), hK9, 11 (four residues), and in hK10 (eight residues). As the residues of the kallikrein loop are in close proximity to the active site, they interact with the substrate. It is believed that the length of the kallikrein loop determines the substrate specificity and catalytic efficiency of these enzymes (Timm, 1997). The crystal structure of human plasma kallikrein (Tang et al., 2005), hK1 (Katz et al., 1998; Laxmikanthan et al., 2005), hK6 (Bernett et al., 2002), and the proenzyme of hK6 (Gomis‐Ruth et al., 2002) have been resolved. The proteolytic activity of kallikreins is regulated in several ways including zymogen activation, endogenous inhibitors such as serpins, and through autolysis leading to inactivation. Dysregulated kallikrein expression is associated with multiple diseases, primarily cancer (Borgono et al., 2004). Several lines of evidence suggest that kallikreins may be involved in cascade reactions and that cross talk may exist with proteases of other catalytic classes.

3.4 Biological Roles of Kallikreins and Their Association with Diseases Plasma kallikrein cleaves the precursor of bradykinin to release the vasoactive peptide (Asakai et al., 1987). hK1 releases lysyl‐bradykinin (kallidin) from low molecular weight kininogen (Clements, 1997). The modulatory effects of the kallikrein–kinin system on the cardiovascular system, particularly in regulating smooth muscle tone and arterial blood pressure and in preventing myocardial ischemia, have also been explored (Marcondes and Antunes, 2005). As hK1 is expressed diversely in different cell types, it was suggested that this enzyme might have additional roles (Schachter, 1979; Bhoola et al., 1992). It has been proposed to be involved in the processing of growth factors and peptide hormones (Mason et al., 1983). Furthermore, hK1 has been shown to process proinsulin, prorenin, procollagenase, angiotensinogen, low‐ density lipoprotein, vasoactive intestinal peptide, and the precursor of vasoactive intestinal peptide (Bhoola et al., 1992). hK2 cleaves seminogelin I and II at different cleavage sites and with lower catalytic efficiency as compared with hK3 (Rittenhouse et al., 1998). hK2 was also shown to activate uPA in vitro (Frenette et al., 1997). hK3 hydrolyses seminogelin I, II, and fibronectin, which leads to the liquefaction of the seminal plasma clot after ejaculation (Lilja, 1985). Other potential substrates of hK3 have also been suggested such as transforming growth factor b (TGF‐b) (Killian et al., 1993), plasminogen (Heidtmann et al., 1999), and insulin‐like growth factor‐binding protein‐3 (IGFBP‐3) (Cohen et al., 1992).

425

426

14

Serine proteases

hK1 is involved in many pathological processes such as inflammation (Clements, 1997), hypertension (Margolius et al., 1974), renal nephritis, and diabetic renal disease (Jaffa et al., 1992). hK7 has been implicated in skin diseases including psoriasis and pathologic keratinization (Sondell et al., 1996; Ekholm and Egelrud, 1999). The relation between cancer and kallikreins is well established (Diamandis and Yousef, 2002; Yousef and Diamandis, 2002a, c; Borgono et al., 2004; Obiezu and Diamandis, 2005). The differential expression of kallikreins is reported in various tumors, (Tanimoto et al., 1999; Dong et al., 2001; Kim et al., 2001; Obiezu et al., 2001; Yousef et al., 2001a, 2002). Recent evidence implicates hKs in many cancer‐related processes, including cell‐growth regulation, angiogenesis, invasion, and metastasis. They have been shown to promote or inhibit neoplastic progression, acting individually or in cascades with other kallikreins and proteases (Borgono and Diamandis, 2004). Kallikreins are widely used serum biomarkers for diagnosis and prognosis of cancer (Diamandis et al., 2000c, d; Luo et al., 2001a, b; Diamandis et al., 2002; Yousef and Diamandis, 2003a; Clements et al., 2004; Obiezu and Diamandis, 2005).

3.5 Potential Roles of Kallikreins in the CNS The potential physiological or pathological roles of kallikreins in the brain was studied mostly in mouse and rat models. hK6 is one of the best characterized enzyme in this aspect. Studies on the mouse ortholog of KLK6 showed that it is expressed in mature oligodendrocytes 2–7 days after maturation, which indicates a role in myelination or turnover of myelin protein (Yamanaka et al., 1999). The rat ortholog of hK6, myelencephalon specific protease (MSP) (Bernett et al., 2002), is specifically expressed in the spinal cord and medulla oblongata and is upregulated in response to glutamate‐receptor mediated cytotoxic injury (Scarisbrick et al., 1997). hK6 has also been implicated in demyelination processes as myelin basic protein (MBP) and myelin oligodendrocyte glycoprotein (MOG) are substrates for this enzyme (Scarisbrick et al., 2002). The mouse genes coding for brain skin serine protease (BSSP) and brain serine protease (BSP) are also suggested to be homologous to KLK6 (Meier et al., 1999; Matsui et al., 2000). hK6 was localized in various tissues using monoclonal and polyclonal antibodies (Petraki et al., 2001). Positivity was observed in the epithelium of the choroid plexus, in the peripheral nerves, in Purkinje cells and the stellate in the cerebellum, but not in the granular cells. Nerve cells showed weak staining throughout the CNS. Studies on MOG‐induced experimental autoimmune encephalomyelitis (EAE), a recognized animal model of multiple sclerosis (MS), showed that EAE caused an increase in the expression of kallikrein 6 mRNA and its protein product throughout the white and gray matter surrounding demyelinating lesions. These results suggest that the differential expression of KLK6 in mature oligodendrocytes and their progenitors is involved in the pathogenesis of MS and EAE (Terayama et al., 2005). Christophi et al. demonstrated tissue‐specific expression patterns and differential regulation of KLK6 in CNS demyelinating disease (Christophi et al., 2004). Terayama et al. reported differential expression of kallikrein 6 and 8 in oligodendrocytes after injury to spinal cord in mice (Terayama et al., 2004). Kallikrein 6 is abundantly expressed by inflammatory cells at sites of CNS inflammation and demyelination in animal models of MS and in human MS lesions. Blaber et al. tested the hypothesis whether kallikrein 6 is a mediator of pathogenesis in CNS inflammatory disease in murine proteolipid protein 139–151‐induced experimental autoimmune encephalomyelitis (PLP139– 151 EAE) (Blaber et al., 2004). They demonstrated that immunization of mice with recombinant hK6 generates antibodies that block hK6 enzymatic activity in vitro, including the breakdown of MBP, and that hK6‐immunized mice exhibit significantly delayed onset and severity of clinical deficits reflected in significantly less spinal cord pathology and meningeal inflammation and in reduced TH1 cellular responses in vivo and in vitro. These data demonstrate for the first time that kallikrein 6 participates in enzymatic cascades mediating CNS progressive inflammatory disorders, including MS. Kallikrein 6 has also been implicated in the pathogenesis of synucleinopathies. a‐Synuclein is an acidic synaptic protein composed of 140 amino acid residues, including 7 incomplete repeats of 11 amino acids and a core sequence of KTKEGV at the N terminus (Maroteaux et al., 1988), which is able to form aggregates under certain circumstances. Accumulation of insoluble a‐synuclein in the brain is observed in Parkinson’s disease (PD), dementia with Lewy bodies, and multiple system atrophy (MSA). Iwata et al. (2003) found that kallikrein

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6 degrades a‐synuclein and colocalizes with pathological inclusions such as Lewy bodies and glial cytoplasmic inclusions. Furthermore, neurosin prevented a‐synuclein polymerization in vitro by reducing the amount of monomer and also by generating fragmented a‐synucleins that themselves inhibited the polymerization. Upon cellular stress, neurosin was released from mitochondria to the cytosol, which resulted in the increase of degraded a‐synuclein species. Downregulation of neurosin caused accumulation of a‐synuclein within cultured cells. These data suggest that kallikrein 6 plays a significant role in physiological a‐synuclein degradation and also in the pathogenesis of synucleinopathies. Mouse neuropsin, the ortholog of hK8, is also a well‐studied kallikrein expressed dominantly in the hippocampal pyramidal neurons. It is suggested to play a role in hippocampal plasticity. Monoclonal antibodies (mAbs) against the enzyme injected intraventricularly reduced the epileptic pattern and inhibited the progression of kindling (Momota et al., 1998). Furthermore, it was shown that kindling induces neuropsin expression (Okabe et al., 1996) and that stimulation of the hippocampus triggers alternative expression (Chen et al., 1995). Neuropsin expression is reduced upon oxidative stress (Akita et al., 1997). It was also indicated that neuropsin might have a role in neuronal development, synaptogenesis (Suzuki et al., 1995), regulation of seizures in kindling (Okabe et al., 1996), and LTP (Komai et al., 2000). Marked abnormalities were also shown in the synapses and neurons in the CA1 subfield of the hippocampus in neuropsin knockout mice, which indicates a role in hippocampal networking (Hirata et al., 2001). Furthermore, it was reported that loss of this protein predisposes to global seizure activity (Davies et al., 1998). The widespread localization and the change of expression pattern of neuropsin during mouse embryonic development indicate that this is an enzyme with multiple functions including neural plasticity, development, and cerebrospinal fluid (CSF) production (Suzuki et al., 1995). Endogenous neuropsin is extracellularly localized in neuronal cell bodies and neurites in mouse hippocampal cell cultures and enhances neurite projection. These observations indicate that neuropsin is involved in neurite outgrowth and fasciculations during development (Oka et al., 2002). Regulatory effect of neuropsin on Schafer collateral LTP in the mouse hippocampus was also shown (Komai et al., 2000). Matsumoto‐Miyai et al. (2003) reported NMDA‐dependent proteolysis of presynaptic adhesion molecule L1 in the hippocampus by neuropsin in mice indicating that kallikrein 8 is involved in NMDA receptor‐dependent synaptic plasticity such as the Schaffer collateral LTP. Xia et al. (2004) examined the potential neuroprotective action of KLK1 gene transfer in cerebral ischemia. Their results indicate that kallikrein gene transfer provides neuroprotection against cerebral ischemia injury by enhancing glial cell survival and migration along with inhibition of apoptosis through suppression of oxidative stress and activation of the Akt–Bcl‐2 signaling pathway.

3.6 Kallikreins in Alzheimer’s Disease The most common form of dementia among older people is AD. It is a neurodegenerative disease sporadic in most of the cases, but genetically determined forms of AD are also known. It has been proposed that the neurodegeneration in AD may be caused by deposition of amyloid‐b peptide (Ab) in plaques in brain tissue, generated from amyloid precursor protein (APP) via proteolysis (Hardy and Selkoe, 2002). According to the amyloid hypothesis, accumulation of Ab in the brain is the primary influence driving AD pathogenesis. The rest of the disease process, including formation of neurofibrillary tangles containing tau protein, is proposed to result from an imbalance between Ab production and Ab clearance. Serine proteases have been proposed to be involved in AD (Selkoe, 1991; Hardy and Selkoe, 2002). hK6, hK8, and hK15 have been reported to be implicated in AD. hK6 has amyloidogenic activity (Magklara et al., 2003), although in cotransfection studies it digested APP695. Immunohistochemical studies showed that hK6 is localized in microvessels and microglial cells in AD brain samples from humans (Yousef et al., 2003), in senile plaques and neurofilament tangles in brains of patients with AD, in Lewy bodies in Parkinson’s disease (Ogawa et al., 2000), and in the epithelial cells of the choroid plexus (Yousef et al., 2003). The expression of KLK6 mRNA in the gray matter is reduced in AD as compared with normal controls, while the concentration of the protein is twofold lower in AD brain tissue extracts compared with normal tissues (Yousef et al., 2003). These results suggest that in AD hK6 may have a role in the degradation of substances such as Ab. The hK6

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levels in CSF and in whole blood are higher in AD compared with controls indicating that this protein is a potential biomarker for the diagnosis and monitoring of AD (Diamandis et al., 2000b). Significantly lower amounts of hK6 in hippocampal and cerebral cortex extracts of patients with AD compared with normal controls were reported (Zarghooni et al., 2002). Diamandis et al. demonstrated significant alterations of hK6, hK7, and hK10 concentrations in CSF of patients with AD and frontotemporal dementia (FTD). hK6, hK7, and hK10 are decreased in the CSF of FTD patients and hK10 is increased in the CSF of patients with AD, in comparison to control subjects (Diamandis et al., 2004). A 12‐fold increase of KLK8 transcription was shown in the hippocampus of patients with AD compared with controls (Shimizu‐Okabe et al., 2001) and an alternatively spliced form of KLK15 mRNAs were isolated from the visual cortex of patients with AD (Dihanich and Spiess, 1994). These data indicate that kallikrein 8 and 15 might also be involved in the pathogenesis of AD.

4

Human Trypsin 4—an Unconventional Serine Protease in the CNS

4.1 Chromosomal Localization and Transcription of Human Trypsin Genes There are several trypsinogen or trypsinogen‐like genes in the human genome of which three (PRSS1, PRSS2, and PRSS3) produce functional trypsins. PRSS1 (coding for human trypsin 1, also called cationic trypsin) and PRSS2 (encoding human trypsin 2, also named as anionic trypsin) are located on chromosome 7q35 intercalated in the b T‐cell receptor (TCR) locus (Rowen et al., 1996). Such organization of the trypsinogen genes has been conserved in mouse (Hood et al., 1995) and chicken (Wang et al., 1995) as well, which may reflect shared functional or regulatory constraints. The PRSS3 gene is localized on chromosome 9p13 and was formed by segmental duplications and translocations originating from chromosomes 7q35 and 11q24 (Rowen et al., 1996, 2005). As a result, PRSS3 transcripts display two variants of exon 1 providing an opportunity for the generation of alternative variants of the protein. Indeed, there are three possible isoforms (isoform A, B, and C) sharing exons 2–5, but differing in their N‐terminal sequences. Isoform C was originally named mesotrypsinogen as its isoelectric point lies between those of human anionic and cationic trypsinogens. This isoform possesses a typical eukaryotic signal sequence characteristic to secreted proteins (a charged residue near the Nterminus followed by hydrophobic and small non-charged amino acids, a b‐turn following the sequence Ala‐ Val‐Ala‐Val, and signal peptidase cleavage site), and it is expressed in the pancreas (Wiegand et al., 1993, Nyaruhucha et al., 1997). In contrast, the other variants are fusion transcripts, called human trypsinogen 4 isoforms A and B lacking a typical signal sequence. These isoforms are expressed in the central nervous system and in epithelial cell lines from prostate, colon, airway, and in normal colonic mucosa (Wiegand et al., 1993, Cottrell et al., 2004). The first exon of trypsinogen 4 is derived from the noncoding first exon of LOC120224, a chromosome 11 gene. LOC120224 codes for a widely conserved transmembrane protein of unknown function. The predicted isoform A might have a highly charged leader peptide consisting of 72 amino acids. Isoform B can be derived from isoform A via alternative translation initiation with a deduced CUG codon initiation site resulting in a shorter, 28 amino acid leader peptide. There are three potential cleavage sites for furin (Arg‐X‐X‐ Arg) in the leader peptide of human trypsinogen 4, opening the door for the processing of the full‐length protein into secreted forms. Indeed, trypsinogen 4 was detected with immunocytochemical methods in vesicles in transfected cell lines and in epithelial cells, and may be secreted from these vesicles (Cottrell et al., 2004). Human trypsin 4 isoforms A and B were expressed in Escherichia coli heterologous system, and the leader peptides were produced by solid phase peptide synthesis in our department. These specimens all showed high affinity toward artificial cardiolipin and phosphatidylcholine membranes in Langmuir balance experiments. Still, the role of these special N‐terminal peptides in the regulation of secretion, expression, trafficking, or localization of these proteins in vivo is yet to be investigated. 50 ‐RACE (rapid amplification of 50 ‐cDNA ends) experiments proved the existence of the mRNA corresponding to isoform B in human brains (also shown by Wiegand et al., 1993) but attempts to isolate the mRNA corresponding to isoform A were unsuccessful. Furthermore, we isolated the expressed protein from postmortem human brain samples by immunoaffinity chromatography using an anti‐28AA leader peptide and an antiprotease domain monoclonal antibody (mAb) as well. N‐terminal amino acid sequencing

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showed that the isolated protein corresponds to isoform B of human trypsinogen 4 (Ne´meth et al., manuscript under revision in FEBS Journal). These data strongly indicate that the predominant (if not the exclusive) variant of this protein is isoform B in the CNS (> Figure 14-4). The immunoreactive material isolated from the temporal cortex using the antiprotease domain antibody contained a minor sequence as well, corresponding to active trypsin. CUG usually codes for leucine, but the generally accepted view is that in cases when it serves as . Figure 14-4 Human trypsin 4 isoforms generated via alternative transcription initiation and alternative splicing. (a): The partial nucleotide and deduced amino acid sequences of exon 1 and the 50 ‐end of exon 2 in the gene encoding human trypsinogen 4. Numbering of the sequence is according to Wiegand et al (1993). Potential furin‐ processing sites are boxed. (b): The sequence of the 50 ‐end of human trypsinogen 4 cDNA as determined by 50 ‐RACE PCR

initiation codon it directs Met‐tRNAimet to the ribosome. However, the sequence analysis of the isolated protein clearly showed that in this case the starting amino acid is a leucine. To confirm this finding, we generated several variants of human trypsinogen 4 DNA templates and transfected HeLa cell lines with the constructs (Ne´meth et al., manuscript in preparation). p72M construct possesses the AUG codon at position 72 (numbering starts from the protease domain) and the CUG codon at position 28 as well, while a truncated construct, named p28L, contained only the CUG codon at position 28. This experimental system allowed for the analysis of alternative transcription initiation with significantly reduced possibility of RNA and protein degradation, which is a serious problem in case of postmortem samples. Western blots together with the N‐terminal amino acid sequences of trypsinogens isolated from the cells transfected with the p72M plasmid imply the expression of both isoforms A and B of human trypsinogen 4. At the same time the expression of only isoform B could be detected in the p28L transfected cells and similarly to brain samples, leucine was identified as the N‐terminal initiator amino acid. Furthermore, to study the expression levels and intracellular localization by confocal fluorescence microscopy, green fluorescent protein (GFP)‐ fusion constructs were also generated. U87 human glioblastoma cells were transiently transfected with the constructs and were immunostained 24 h posttransfection with mAbs raised against the protease domain or against the 28 amino acid leader peptide. The GFP reporter protein always colocalized with the immunostaining with both antibodies indicating that trypsinogen 4 may exist mainly in an inactive zymogen form in the cytosol. The relative expression level was always higher when translation was initiated from an AUG codon. The relative expression level of isoform B was highly dependent on the initiator codon, but was relatively independent of the length of the 50 ‐untranslated region of the recombinant mRNA.

4.2 Regional Distribution of Human Trypsin 4 mRNA and Protein in the Brain We also examined the distribution of human trypsinogen 4 at mRNA and protein levels in 17 different areas of the human brain (Siklo´di et al., unpublished results). The relative amount of trypsinogen 4 mRNA was measured using quantitative real‐time PCR (> Figure 14-5). The quantity of human trypsinogen 4 mRNA

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. Figure 14-5 Relative quantity of human trypsinogen 4 mRNA in different brain regions. Transcription levels were measured with quantitative real‐time PCR and normalized to the amount of b‐actin mRNA

relative to β-actin mRNA is in the 3.0–7.4% range. Evaluating the data with analysis of variance showed that the relative quantity of human trypsinogen 4 mRNA is significantly different in the examined brain areas. High relative amounts were detected in cerebellar cortex, occipital cortex, parahippocampal cortex and periaqueductal gray matter samples. Low relative quantities were found in cortical white matter and plexus choroideus samples, and intermediate amounts could be observed in the rest of the regions. The greatest (24-fold) difference is observed between the cortical white matter and the cerebellar cortex samples. We determined the distribution of the expressed protein in different brain areas using sandwich ELISA combined with immunoprecipitation (> Figure 14-6). Both antiprotease domain and anti‐28AA leader peptide mAbs were applied to capture the antigens from the brain extracts by immunoprecipitation, which enabled us to measure the quantity of the zymogen and the active enzyme separately. The amount of total protein was in the range of 10–30 ng/g wet tissue. In seven areas (frontopolar cortex, occipital cortex, hypothalamus, putamen, thalamus, periaqueductal gray matter, and cervical spinal cord), the total amount (zymogen þ active enzyme) was roughly equal to the amount of the zymogen, while in the rest of the areas (bulbus olfactorius, cingulate cortex, somatomotor cortex, cortical white matter, hippocampus, substantia nigra, pons, cerebellar cortex, cerebellar white matter, and medulla oblongata) the amount of the zymogen

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. Figure 14-6 The amount of human trypsinogen and trypsin 4 in different areas of the brain. Immunoprecipitation was carried out with antiprotease domain (shaded columns) or anti‐28AA leader peptide (white columns) capture monoclonal antibodies

was substantially less than the total amount. From these data we assume that the extent of activation depends on brain areas. The highest degree of activation was found in cerebellar cortex, medulla oblongata, and hippocampus. The relatively high amount of trypsinogen and/or trypsin 4 in the cerebral and cerebellar white matters is indicative of the expression of this protease in glial elements. Indeed, immunohistochemical studies showed that the protein is localized in glial cells and neuronal perikarya (Gallatz et al., unpublished results). Human trypsinogen‐4‐like immunoreactivity was localized in human brain samples using both antiprotease domain and anti‐28AA leader peptide mAbs. In the spinal cord, glial cells showed strong immunoreactivity in the funiculi. The distribution, shape, and size of the immunostained cells were consistent with those of GFAP (glial fibrillar acidic protein, an astrocyte marker)‐stained alternate sections indicating that the immunopositive cells in the spinal white matter are most probably astrocytes. Neuronal cells in the ventral horn and the central gray matter were also immunostained for human trypsinogen 4 but not for GFAP. Human trypsinogen‐4‐like immunopositive glial cells and fibers were observed in the cortical white matter (frontal cortex) and in neuronal cells in cortical layers III, IV, and V. Similar to the spinal cord, glial cells were equally stained with antihuman trypsinogen 4 and anti‐GFAP antibodies but not with anti‐28AA leader peptide mAb. The low to moderate levels of homogenous and uncharacterized human trypsinogen‐ 4‐like immunostaining in the gray matter of the cerebral cortex and in the granule cell layers of the cerebellum may suggest the presence of this protease in the neuronal matrix, i.e., in the extracellular space.

4.3 Structural and Biochemical Characterization of Human Trypsin 4 Human trypsinogen 4 was first isolated from pancreatic fluid (Rinderknecht et al., 1984). The active enzyme was characterized biochemically and was found to be resistant to canonical and serpin‐type inhibitors as well. On the basis of sequence analysis of the cloned cDNA coding for human trypsin 4, it

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has been shown that the enzyme possesses most of the major features characteristic for the trypsin‐like serine protease subfamily (Wiegand et al., 1993). The polyanionic cluster followed by the activation cleavage site, the catalytic triad (His57, Asp102, Ser195), and an aspartate at position 189 and glycine residues at positions 212 and 222 that determine the properties of the substrate‐binding pocket are typical features among trypsin‐like proteases. However, exceptional properties of this enzyme were also revealed: it encompasses only ten conserved cysteine residues instead of twelve and possesses an arginine residue at position 193 (S20 subsite according to the Schechter–Berger nomenclature (Schechter and Berger, 1967)) in place of the highly conserved glycine. The resolution of the X‐ray structure of human trypsin 4 complexed with the inhibitor benzamidine (Katona et al., 2002) revealed the basis of the unusual features of this protease besides characters typical to conventional (pancreatic) trypsins. The overall fold of the enzyme was found to be highly similar to human trypsin 1; the deviation of the Ca positions is only 0.5 a˚ throughout the molecule. Kinetic analysis of the hydrolysis of small synthetic substrates showed that human trypsin 4 has a similar catalytic constant (kcat) value with a slightly lower Michaelis constant (Km) value when compared with bovine trypsin yielding a slightly higher catalytic efficiency (kcat/Km) (Katona et al., 2002). We compared the catalytic efficiency of the wild type and the R193G mutant human trypsin 4 on small synthetic substrates (Z‐Gly‐Pro‐Arg‐4‐nitroanilide and PyrGlu‐Gly‐Arg‐7‐amido‐4‐methylcoumarin) that do not possess P20 residues that would interfere with Arg193. Our data showed that the substitution of Gly193 with arginine results in the deceleration of acylation and acceleration of the deacylation step. Both kcat and Km values were slightly increased in the wild‐type enzyme, while the catalytic efficiency (kcat/Km) was not significantly altered. Two distinct hypotheses might arise to account for these data: (1) the specific geometry of the catalytic triad, the oxyanion hole, and the subsites S1–S4 are not perturbed by the mutation or (2) though the structure of the S1 site and the oxyanion hole are distorted proper conformation can be restored upon occupancy of the S1 site. This latter theory was confirmed by homology modeling and kinetic data as well in a similar case, where the position 193 is also occupied by a nonglycine residue (Schmidt et al., 2004). Faster rate of carbamylation of NH2‐Ile16, slower rate inhibition by diisopropyl fluorophosphate, and weaker affinity toward an S1‐site probe indicated that the unoccupied active site of the Gly193Glu mutant factor XI is distorted. It was also presumed and confirmed by homology modeling that the conformation of the 192–193 peptide bond is altered in the active enzyme, and thus the amide nitrogen of Glu193 does not point correctly toward the oxyanion hole. Such an argument might hold for human trypsin 4 as well. To study rapid transient kinetics and thermodynamic consequences of a nonglycine residue at position 193, we expressed wild type and R193G mutant human trypsin 4 and monitored their reactions on 4‐methylumbelliferyl 4‐guanidinobenzoate (MUGB) using transient kinetic methods (To´th et al., 2006.). Acylation is slightly (3–4 times) slower in the wild‐type enzyme. The acyl‐enzyme complex breaks down for both enzymes, and deacylation is slightly (5–6 times) faster by the wild‐type enzyme than by the R193G mutant. The formation of the first tetrahedral intermediate proved to be highly exothermic and was accompanied by a large entropy decrease for the wild‐type enzyme, while it is endothermic compensated with entropy gain for the R193G mutant. This difference in the energetic profiles indicates much more extended structural and dynamic rearrangements during the formation of the first tetrahedral intermediate in wild‐type human trypsin 4 than in the R193G mutant enzyme, which may contribute to the biological function of this protease. The presence of arginine at position 193 may induce strain in the backbone of the active form of trypsin, which might account for the difference in the thermodynamics of the tetrahedral intermediate formation. We suppose that a nonglycine residue at this position does not cause significant alterations in the structure of the active enzyme, but evokes mechanical stress that can be revealed by thermodynamical analysis. Furthermore, this presumption is indicated by our preliminary experiments in which the rate constant of the structural change from an inactive to the active conformation of wild‐type human trypsin 4 was smaller than that of the R193G mutant in pH‐jump transient kinetic studies. Since there is a large rotation around position 193 during the pH‐jump activation, the smaller rate constant indicates the presence of a structural or mechanical stress in the 192–194 backbone (To´th et al., manuscript under review in proteins). Analysis of the crystal structure also revealed that the bulky side chain of Arg193 occupies an extended conformation, and thus fills the S20 subsite and contributes to the particular clustering of positive charges

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around the primary specificity pocket as well. These unusual features account for the inhibitor resistance toward canonical inhibitors, the inability to activate human chymotrypsinogen (Rinderknecht et al., 1984), cationic and anionic human trypsinogen (Szila´gyi et al., 2001), and restricted substrate specificity toward macromolecular substrates. In a recent study it has been shown that human trypsin 4 readily cleaves the reactive‐site peptide bond of soybean trypsin inhibitor (STI) and degrades human pancreatic secretory inhibitor (hPSTI), and the role of Arg193 in these interactions has been further confirmed by site‐directed mutagenesis (Szmola et al., 2003). The substitution of Arg193 with a glycine residue (R193G) reverts these unusual phenomena and converts the enzyme to a conventional trypsin. Human trypsin 4 was shown to be resistant to STI, bovine pancreatic trypsin inhibitor (BPTI), and hPSTI (Nyaruhucha et al., 1997). In addition, the importance of the unique amino acid substitution at position 193 was first emphasized: it has been presumed that a positively charged residue at this position might interfere with trypsin inhibitors and macromolecular substrates upon interaction. To probe this hypothesis, we analyzed the hydrolysis of the peptide Z‐Gly‐Pro‐Arg‐Met‐Gly‐Phe‐NH2 by wild‐type human trypsin 4 and human trypsin 1 by following the product generation using HPLC and by applying the peptide as a ‘‘silent substrate’’ in a competitive system with the signal‐generating substrate Z‐Gly‐Pro‐Arg‐pNA. The resulting progress curves were analyzed by global fitting. Our data show that the substitution of Gly193 by Arg results in a slight increase of kcat, while Km drops an order of magnitude, which is probably caused by steric interference with substrate binding. We also examined the effect of this mutation on the hydrolysis of a polypeptide‐type substrate. Chymotryptic myosin subfragment‐1 from rabbit skeletal muscle was digested by wild‐type human trypsin 4 and human trypsin 1 at a ratio of 1:100 and 1:5000, respectively. Samples taken at different periods were run on SDS‐polyacrylamide gels. The intensity of the 95‐kDa band was measured by densitometry, normalized to its initial density, and plotted against time. By fitting single exponential decay functions to the data, the value of kcat/Km can be calculated from the exponent of the function. Our results indicate that human trypsin 4 digests myosin subfragment‐1 200‐times less effectively than human trypsin 1. Taking these observations together, we conclude that substitution of Gly193 by an arginine results in the perturbation of the H‐bonding interactions in the oxyanion hole. This results in the destabilization of the transition‐state complex, thus decreasing the rate of acylation and increasing the rate of deacylation, but the catalytic efficiency is only slightly affected. In case of peptide and protein substrates, the binding is also influenced by the substitution. A bulky and charged residue in the S20 subsite sterically interferes with the binding of the substrate to the enzyme surface, thus increasing the value of Km.

4.4 On the Potential Role of Human Trypsin 4 in Physiological or Pathological Processes in the CNS In search for the function of this unconventional protease in the CNS, experiments have been made in several diversified fields. Protease‐activated receptors (PARs) are a family of four G‐protein‐coupled receptors (GPCRs) triggering signal transduction pathways upon being cleaved by serine proteases (Macfarlane et al., 2001; Ossovskaya and Bunnett, 2004). The tethered ligand domains become exposed upon cleavage and bind to the cleaved receptors and are thus activated. PARs play a role in the regulation of hemostasis, inflammation, pain, and healing. Epithelial cells lines from prostate, colon, and airway and human colonic mucosa were shown to express mRNA encoding PAR‐2, trypsinogen 4, and enteropeptidase that activate the zymogen (Cottrell et al., 2004). Exposure of cell lines expressing PAR‐2 and PAR‐4 to trypsin 4 increased the intracellular Ca2þ concentration and strongly desensitized cells to PAR agonists, whereas there were no responses in cells lacking these receptors. From these data, Cottrell et al. concluded that human trypsin 4 is a potential agonist of PAR‐2 and PAR‐4 in epithelial tissues and its resistance to endogenous trypsin inhibitors may permit prolonged signaling. However, in a more recent study Grishina et al. (2005) found that human mesotrypsin did not induce a PAR‐mediated Ca2þ response in human epithelial cells (HBE (human bronchial epithelial), A549 (human pulmonary epithelial), and HEK (human embryonic kidney)‐293 cells expressing functional PAR‐1 and PAR‐2) even at high concentrations. In addition, mesotrypsin did not affect the magnitude of PAR activation by subsequently added bovine

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trypsin. They also investigated human astrocytoma 1321N1 cells, which express PAR‐1 and some PAR‐3, but no PAR‐2. High concentrations of mesotrypsin produced a relatively weak Ca2þ signal, apparently through PAR‐1 activation. Mesotrypsin, in contrast to cationic and anionic trypsin, cannot activate or disable PARs in human epithelial cells, demonstrating that the receptors are not substrates for this isoenzyme. On the other hand, mesotrypsin activates PAR‐1 in human astrocytoma cells. From these data it has been assumed that human trypsin 4 might play a role in protection/degeneration or plasticity processes in the human brain. PRSS3 encoding human trypsinogen 4 has also been implicated as a putative tumor suppressor gene (Marsit et al., 2005.) and its expression was also reported to be associated with metastasis (Diederichs et al., 2004). In order to test whether trypsinogen 4 is involved in CNS diseases or injury response in mammalian brains, a mouse model was developed expressing human trypsinogen 4 specifically in neurons (Minn et al., 1998). Immunocytochemical analysis of the brains of transgenic mice revealed a striking enhancement of GFAP expression in astrocytes. This remarkable astrocytic reaction was detected in the brains of mice as young as 2 months and did not diminish in the older animals tested. However, gross evidence for neurodegeneration or for reactive microglial cells was not found. In addition, the possible participation of trypsin 4 in the metabolism of the amyloid precursor protein (APP) was investigated by immunostaining brains from transgenic mice with Ab (bA4) antibodies. Immunocytochemical staining of brains of 1‐year‐ old transgenic mice revealed an intense intracellular bA4‐like signal in neurons. Mechanisms regulating neuronal migration during development are still unclear. Extracellular matrix cues, target site released factors, and components of the migratory neurons themselves are all playing roles in directing neurons to their appropriate locations. Drapkin et al. have studied the effects of proteases and their inhibitors on the extracellular matrix and the consequences to the migration of gonadotropin‐ releasing hormone (GnRH) neurons in the embryonic chick (Drapkin et al., 2002). Chick GnRH neurons differentiate in the olfactory epithelium, migrate along the olfactory nerve, and enter the forebrain. Affigel blue beads were used to deliver a serpin‐type serine protease inhibitor, protease nexin‐1 (PN‐1, also called glia‐derived nexin (GDN)), and a target protease, trypsin, to the olfactory epithelium coincident with initiation of GnRH neuronal migration. PN‐1 inhibited neuronal migration while trypsin accelerated their transit into the CNS. Prior to initiation of migration, neither PN‐1 nor trypsin altered the timing of neuronal exit. Trypsin did, however, accelerate the timing of neuronal crossing into the nerve–forebrain junction. These data support the hypothesis that protease activity modulates neuronal movements across barriers. Moreover, the data suggest, for the first time, that aspects of GnRH neuronal migration may be cell autonomous but modulated by extracellular matrix alterations. To shed light on the function of human trypsin 4 in the CNS, we searched for potential natural substrates. Human brain extracts were either incubated with or without human trypsin 4, and then separated using two‐dimensional SDS‐polyacrylamide gel electrophoresis. The spots whose quantity or position on the gel changed upon the human trypsin 4 treatment were excised from the gel and identified using matrix‐assisted laser desorption ionization mass spectrometry (MALDI‐MS) following tryptic digestion. Several proteins (a B‐crystallin, lactate dehydrogenase B, isocitrate dehydrogenase, septin 5, creatin kinase B, cofilin 1, collapsin response mediator protein 2, capping protein a 2, and a tubulin) are degraded in the brain protein extract upon treating with human trypsin 4. Five out of the nine identified potential substrate proteins are involved in the synthesis and regulation of the cytoskeletal system (P. Medveczky, E. Siklo´di, J. To´th, M. Palkovits, L. Szila´gyi and L. Gra´f, unpublished results). Demyelination, the breakdown of the major membrane protein of the CNS, is involved in many neurodegenerative diseases; proteases participating in this process are thus potential targets of therapy. In our in vitro study the proteolytic actions of calpain, human trypsin 1, and human trypsin 4 were compared on lipid‐bound and free human MBP as substrates (Medveczky et al., 2006). Both calpain and human trypsin 4 are expressed in human brains, and thus their peptidase activities may be of some physiological or pathological relevance. The generated fragments were identified using N‐terminal amino acid sequencing and mass spectrometry. Analysis of the degradation products showed that human trypsin 4 selectively cleaves the Arg80‐Thr81 and Arg98‐Thr99 peptide bonds in the lipid bound form of human MBP. On the basis of this data we synthesized a peptide segment consisting of amino acids 94–104 of MBP

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(IVTPRTPPPSQ) that contains the specific trypsin 4 cleavage site Arg98‐Thr99. In vitro studies on the hydrolysis of this synthetic peptide by trypsin 4 confirmed the above results. These findings are of biological interest as the major autoantibodies found in patients with MS recognize the peptide segment 80–96 of MBP. Our results suggest that human trypsin 4 may be one of the candidate proteases involved in the pathomechanism of MS.

5

Concluding Remarks

Serine proteases are a group of proteolytic enzymes in which the catalytic mechanism depends upon the hydroxyl group of the catalytic serine residue. Enzymes belonging to this family of proteases are highly diversified and widespread throughout organisms. The genes encoding these proteases are clustered in the genome as they most probably arose from a common ancestor via gene duplication and fusion events during evolution. Many biological processes that require specific and limited proteolysis are mediated by serine proteases. Such enzymatic activities are blood clotting and fibrinolysis, inflammatory responses, tissue remodeling, programmed cell death, and processing hormones, growth factors, and receptors. There is an increasing body of evidence showing that serine proteases are widely expressed and fulfill diverse roles in the CNS. Serine proteases contribute to structural plasticity associated with learning and memory, neurite outgrowth, regulation of neuronal survival, synaptogenesis, myelin turnover, and development. These proteases have also been implicated in the pathophysiology of neurodegenerative disorders such as AD, PD, and MS. The activities of these enzymes must be under strict regulation through transcription, translation, autolysis, and interaction with natural inhibitors. A fine‐tuned balance of proteolysis is required in the CNS to maintain normal physiology; perturbation of this equilibrium may lead to serious disturbances in neuronal function. The expression of diverse serine proteases in the CNS possessing different specificity reflects the complex regulatory mechanisms of the CNS and also provides opportunities to fine‐ tune these physiological and pathological processes.

Acknowledgments This study was supported by a Hungarian Scientific Research Fund grant OTKA to L. Gra´f (T047154, TS 049812), L. Szila´gyi (T037568), and to J. Gergely (TS 0044711). The huge volume of literature in the field of serine proteases made it impossible to cite every study in this review. As such, the authors would like to apologize to their colleagues for any unintentional oversight or omission of their studies.

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Calpain as a Target for Prevention of Neuronal Death in Injuries and Diseases of the Central Nervous System

S. K. Ray . M. K. Guyton . E. A. Sribnick . N. L. Banik

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447

2 2.1 2.1.1 2.1.2 2.2 2.2.1 2.2.2 2.3 2.4 2.5

Increase in Intracellular Free Ca2þ and Calpain Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 448 Ligand‐Gated Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 448 Glutamate Receptor Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 448 Ligand‐Gated Ca2þ Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 449 Voltage‐Gated Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 450 Voltage‐Gated Ca2þ Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 450 Reverse Naþ/Ca2þ Exchanger . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 450 Ca2þ Extrusion and Sequestration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 450 Ca2þ‐Binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 Calpain Activation and Therapeutic Opportunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451

3 3.1 3.1.1 3.1.2 3.2 3.2.1 3.2.2 3.3 3.3.1 3.3.2

Calpain Involvement in Neuronal Death in CNS Injuries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 Ischemic Brain Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 Necrotic and Apoptotic Neuronal Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 Calpain Activation in Neuronal Death and Calpain Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452 Spinal Cord Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452 Ca2þ Accumulation, Calpain Activation, and Neuronal Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452 Calpain Inhibition for Neuroprotection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453 Traumatic Brain Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453 Changes in Ca2þ Homeostasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453 Calpain Activation and Calpain Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453

4 4.1 4.1.1 4.1.2 4.2 4.2.1 4.2.2 4.3 4.3.1 4.3.2 4.4 4.4.1 4.4.2 4.5

Calpain Involvement in Neuronal Death in CNS Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Role of Calpain in Neurodegeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Calpain Inhibition as a Therapeutic Strategy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 455 Amyotrophic Lateral Sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 455 Mechanism of Motoneuron Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 455 Calpain Activation in Motoneuron Death and Calpain Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 455 Epileptic Seizures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 456 Excessive Intracellular Free Ca2þ and Calpain in Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 456 Oxidative Stress Leading to Calpain Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 456 Huntington’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 457 Mechanism of Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 457 Excitotoxicity, Ca2þ Influx, and Calpain Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 457 Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 457

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Calpain as a target for prevention of neuronal death in injuries and diseases of the CNS

4.5.1 4.5.2 4.6 4.6.1 4.6.2

a‐Synuclein and Oxidative Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Calpain Activation and Calpain as a Target for Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Multiple Sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Multiple Mechanisms of Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Potential Role of Calpain in Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 459

5

Conclusion and Future Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 459


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Abstract: Multiple mechanisms are known to contribute to an increase in the intracellular free Ca2þ level. Uncontrolled upregulation of the intracellular free Ca2þ level has been implicated in the pathogenesis of injuries and diseases of the central nervous system (CNS). An increase in the intracellular free Ca2þ level activates the Ca2þ‐dependent cysteine protease calpain that can mediate neuronal death. Various studies demonstrated that calpain could degrade key cellular substrates so as to impair structural integrity and normal cellular function, leading to both necrotic and apoptotic neuronal death in many CNS disorders. It is now highly recognized from cell culture and animal model studies that calpain has an important role in the mediation of neuronal death in CNS injuries such as ischemic brain injury, spinal cord injury (SCI), and traumatic brain injury (TBI), and also diseases such as Alzheimer’s disease (AD), amyotrophic lateral sclerosis (ALS), epileptic seizures, Huntington’s disease (HD), Parkinson’s disease (PD), and multiple sclerosis (MS). In many of these cases, use of calpain inhibitors provided neuroprotection suggesting that calpain could be a potential target for therapeutic interventions. However, currently available calpain inhibitors lack sufficient drug properties that should be addressed adequately before their clinical applications. Importantly, many investigators are now taking interest in developing clinically suitable calpain inhibitors for functional neuroprotection in CNS injuries and diseases. List of Abbreviations: AD, Alzheimer’s disease; ALS, amyotrophic lateral sclerosis; AMPAR, a‐amino‐3‐ hydroxy‐5‐methyl‐4‐isoxazole propionate receptor; CaBPs, Ca2þ‐binding proteins; CANP, Ca2þ‐activated neutral protease; CNS, central nervous system; EAA, excitatory amino acid; EAE, experimental allergic encephalomyelitis; ER, endoplasmic reticulum; HD, Huntington’s disease; iGluR, ionotropic glutamate receptor; IP3, inositol triphosphate; IP3RP3 receptors; KAR, kainate receptor; LGCC, ligand‐gated Ca2þ channels; LGIC, ligand‐gated ion channels; MAP2, microtubule‐associated protein 2; MBP, myelin basic protein; mGluR, metabotropic glutamate receptor; MS, multiple sclerosis; NFP, neurofilament protein; NMDAR, N‐methyl‐D‐aspartate receptor; ORP150, oxygen‐regulated protein‐150 kDa; PD, Parkinson’s disease; PIP2, phosphoinositol biphosphate; PLA2, phospholipase A2; ROS, reactive oxygen species; RyR, ryanodine receptors; SCI, spinal cord injury; TBI, traumatic brain injury; VGCC, voltage‐gated Ca2þ channels; VGIC, voltage‐gated ion channels

1

Introduction

Calpain is a calcium (Ca2þ)‐dependent cysteine protease localized and activated by autolysis in the cytosol of all cells; it is also translocated to membranes and activated following interaction with the membrane‐ bound phospholipids (Croall and DeMartino, 1991; Ray and Banik, 2002). This Ca2þ‐activated neutral protease (CANP) was first identified in rat brains (Guroff, 1964) and muscles (Meyer et al., 1964). The old name CANP has currently been replaced by ‘‘calpain’’ that stands for calcium‐activated papain‐like cysteine protease. Many calpain subunits and their homologs have recently been identified by cDNA cloning. During 1995–2005, research has grown remarkably on calpain involvement in the pathophysiology of human diseases, resulting in the discovery of different members of the calpain superfamily (http://www.ag.arizona. edu/calpains). In general, the term ‘‘calpain’’ refers to ubiquitously expressed calpain 1 and calpain 2. Obviously, some other calpain homologs are not expressed ubiquitously but predominantly in a limited number of mammalian tissues. Therefore, on the basis of the tissue distributions, calpains can be broadly classified as ubiquitous and tissue-specific (Sorimachi et al., 1997). The tissue‐specific calpains may have functions in those tissues in which they are expressed predominantly. It should be noted that currently we know very little about the tissue‐specific calpains. The ubiquitous expression of calpain 1 and calpain 2 in mammals strongly suggests their involvement in some important and fundamental cellular functions. Although the precise physiological functions of calpain remain mostly unknown, it is considered to participate in a number of cellular signaling pathways that are regulated by intracellular free Ca2þ concentrations (Carafoli and Molinari, 1998; Sato and Kawashima, 2001). The precise physiological function of calpain will remain mostly unknown until we can determine the exact mechanism that controls the in vivo Ca2þ threshold for calpain activation. Various studies suggest that in vivo Ca2þ threshold for calpain activation may drop to a substantially low level

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following translocation and interaction of calpain with certain phospholipids on the membrane. However, when intracellular free Ca2þ levels exceed this threshold, uncontrolled and prolonged calpain activation can occur leading to the development of pathophysiological processes in the central nervous system (CNS). Increased activation of ubiquitous calpain has been implicated in the mediation of neurodegeneration in various CNS injuries such as ischemic brain injury, spinal cord injury (SCI), and traumatic brain injury (TBI), and CNS diseases such as Alzheimer’s disease (AD), amyotrophic lateral sclerosis (ALS), epileptic seizure, Huntington’s disease (HD), Parkinson’s disease (PD), and multiple sclerosis (MS) (Bartus et al., 1995b; Banik and Shields, 1999; Stracher, 1999; Vanderklish and Bahr, 2000; Huang and Wang, 2001; Ray et al., 2001c, 2003). Because calpain has been found to be involved in the mediation of neuronal death in various CNS injuries and diseases, there is an increasing interest in developing and using calpain inhibitors for functional neuroprotection.

2

Increase in Intracellular Free Ca2þ and Calpain Activation

Calpain activation is absolutely dependent on an increase in intracellular free Ca2þ levels (Croall and DeMartino, 1991; Ray and Banik, 2002). Normal neuronal function requires a well‐regulated intracellular Ca2þ homeostasis (Miller, 1987; Clapham, 1995), which is disrupted following induction of cell death by various stress stimuli (Nicotera et al., 1992). Multiple mechanisms currently known to contribute to the intracellular Ca2þ overload include Ca2þ entry through ligand‐gated ion channels (LGIC) and voltage‐ gated ion channels (VGIC), Ca2þ release from intracellular storage organelles, and impairment of Ca2þ extrusion and sequestration due to failure of energy‐dependent mechanisms (> Figure 15‐1).

2.1 Ligand‐Gated Ion Channels The ligand‐gated ion channels (LGIC) superfamily of receptors has an intrinsic ion channel that undergoes conformational change upon binding of an extracellular ligand to its site, allowing the opening of the ion pore (Barnard, 1992). This gating mechanism ceases to exist if the ligand leaves the binding site. All members of LGIC are usually composed of a pentameric arrangement of subunits surrounding a central ion pore. The subunits of the LIGC are needed for assembly, structural integrity, ligand binding, and ion permeation.

2.1.1 Glutamate Receptor Channels An increased level of an excitatory amino acid (EAA) is potentially harmful to neurons. The main EAA in the CNS is glutamate that may stimulate ionotropic glutamate receptor (iGluR) channels and metabotropic glutamate receptor (mGluR) channels (Mayer and Armstrong, 2004; Nagy et al., 2004). The iGluR superfamily consists of three subtypes: N‐methyl‐D‐aspartate receptor (NMDAR), a‐amino‐3‐hydroxy‐5‐ methyl‐4‐isoxazole propionate receptor (AMPAR), and kainate receptor (KAR) that vary in molecular structure and degree of permeability to different ions (Dingledine et al., 1999). Over stimulation of iGluR channels by glutamate causes excitotoxicity and Ca2þ influx that plays an important role in neuronal death in many CNS disorders (Nagy et al., 2004). Individual studies indicate that activation of NMDAR (Ahmed et al., 2002) and AMPAR (Bennett et al., 1996) facilitates Ca2þ influx, leading to neurodegeneration. Comparative studies suggest that the NMDAR channels play the predominant role for Ca2þ influx in neurons under stress (Schneggenburger et al., 1993; Rogers and Dani, 1995). Glutamate not only stimulates NMDAR channels but also interacts with mGluR channels coupled to G‐proteins for the hydrolysis of membrane phosphoinositol biphosphate (PIP2) and for the intracellular production of inositol triphosphate (IP3) that binds to IP3 receptors (IP3R) on endoplasmic reticulum (ER), releasing Ca2þ into the


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. Figure 15‐1 Multiple mechanisms collectively contribute to a sustained increase in intracellular free Ca2þ, leading to calpain activation and neuronal death. Ligand‐gated ion channels (e.g., NMDAR) are the primary sources of Ca2þ influx. Voltage‐gated ion channels (e.g., VGCC) are the secondary sources of Ca2þ influx. Activation of metabotropic glutamate receptors (mGluR) may indirectly contribute to increased intracellular free Ca2þ levels. Extrusion of Ca2þ from neurons via energy‐dependent Ca2þ pump (Ca2þ‐ATPase) fails due to an abrupt fall in ATP levels following CNS stress. Intracellular Naþ concentration rises with membrane depolarization and a ubiquitous Ca2þ transport mechanism works as a reverse Naþ/Ca2þ exchanger to export Naþ and import Ca2þ into the neurons. Sequestration of Ca2þ in endoplasmic reticulum (ER) and mitochondria are disrupted following CNS injuries and diseases. Therefore, sustained increase in intraneuronal free Ca2þ causes calpain activation, which degrades different classes of proteins and affects signal transduction mechanisms. Cleavage of calpain substrates and impairment of cellular signal transduction processes lead to neuronal death

cytosol (Berridge, 1993). It is now well established that activation of mGluR causes release of Ca2þ from the intracellular organelles (Nash et al., 2001).

2.1.2 Ligand‐Gated Ca2þ Channels Ligand‐gated Ca2þ channels (LGCC) are involved in releasing Ca2þ from the intracellular stores so as to regulate proliferation, exocytosis, gene expression, and apoptosis. These channels may also be involved in pathological conditions. They are known to play critical roles for an increase in intracellular free Ca2þ concentration in neuronal cells (Bertolino and Llinas, 1992). The two major families of Ca2þ‐releasing channels are ryanodine receptors (RyR) and IP3 receptors (IP3R), which are found on intracellular Ca2þ storage/release organelles (MacKrill, 1999; Fill and Copello, 2002). Ryanodine binds to RyR and triggers Ca2þ release by locking the Ca2þ channel in an open configuration. There are three subtypes of RyR (RyR1, RyR2, and RyR3) and also three subtypes of IP3R (IP3R1, IP3R2, and IP3R3) with their expression in

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distinct tissues and cells. Stimulation of RyR by increased cytosolic Ca2þ with no requirement of IP3 is a plausible mechanism for releasing Ca2þ from the ER (Berridge, 1993; Charles et al., 1993; Ehrlich, 1995). Cytosolic Ca2þ also regulates IP3R channels that require the presence of IP3 for functional activity.

2.2 Voltage‐Gated Ion Channels The members of the voltage‐gated ion channels (VGIC) superfamily including voltage‐gated Ca2þ channels (VGCC) remain closed at resting membrane potential; however, alteration in membrane potential induces conformational changes that result in opening of the ion pore (voltage‐dependent activation) to allow the ion to enter the cell (Ashcroft, 2000). In a resting cell, the membrane is polarized and the extracellular part of the pore is closed while the intracellular part of the gating mechanism remains open. Upon activation of the cell, the membrane depolarizes, opening the extracellular part of the pore to allow the ion to enter the cell.

2.2.1 Voltage‐Gated Ca2þ Channels Voltage‐gated Ca2þ channels play an important role in the regulation of cell functions such as motility, gene expression, cell cycle, and cell death. They are the major routes of Ca2þ translocation across the plasma membrane of excitable cells. These channels can be crucial to increase intracellular free Ca2þ concentration in neuronal cells (Bertolino and Llinas, 1992). In the course of CNS injuries and diseases, failure of energy‐ dependent ionic pumps (e.g., Naþ/Kþ‐ATPase) causes breakdown of transmembrane ionic gradients (Lees, 1991), allowing extracellular Ca2þ to enter into the neurons via opening of VGCC. Currently, there are six classes of VGCC, which are distinguished depending on their sensitivity to high (L‐type, N‐type, P‐type, Q‐type, and R‐type channels) and low (T‐type channels) voltage changes (Triggle, 1994; Arikkath and Campbell, 2003). Strong membrane depolarizations induce almost continuous Ca2þ influx through L‐type channels, whereas weak depolarizations stimulate T‐type channels for transient and tiny Ca2þ currents (Nowycki et al., 1985; Snutch and Reiner, 1992; McClesky, 1994).

2.2.2 Reverse Naþ/Ca2þ Exchanger After the failure of energy‐dependent processes, another potentially important mechanism of Ca2þ influx is the Naþ/Ca2þ exchanger (Stys, 1998). Following energy failure, transmembrane Naþ gradients collapse and the Naþ/Ca2þ exchanger acts as a reverse Naþ/Ca2þ exchanger to pump Ca2þ in and Naþ out instead of pumping Ca2þ out and Naþ in cells (White and Reynolds, 1995). The operation of the reverse Naþ/Ca2þ exchanger may be the primary mechanism of Ca2þ influx in CNS white matter that lacks glutamate receptors (Stys et al., 1992).

2.3 Ca2þ Extrusion and Sequestration Energy depletion also directly impairs ATP‐dependent Ca2þ extrusion and sequestration. Extrusion of Ca2þ from the cells is slowed down directly due to the failure of the plasma membrane Ca2þ pump (i.e., Ca2þ‐ATPase) that acts in a ATP‐dependent fashion (Carafoli, 1987) and also indirectly due to operation of the reverse Naþ/Ca2þ exchanger as mentioned above. Sequestration of Ca2þ in the ER is rather a slow process that alone may not bring down cytosolic Ca2þ level at the same rate at which Ca2þ enters into the cells after stimulation (Nachshen, 1985). Mitochondrial sequestration of Ca2þ occurs when intracellular free Ca2þ rises to micromolar levels (Blaustein, 1988; Gunter and Pfeiffer, 1990). However, excessive mitochondrial sequestration of Ca2þ may contribute to cytosolic acidosis (Hartley and Dubinsky, 1993) and free radical formation (Reynolds and Hastings, 1995), forcing the mitochondria to be essentially indolent.


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2.4 Ca2þ‐Binding Proteins Interestingly, more than 99% of the Ca2þ that enters into the neurons does not stay as free Ca2þ in the cytoplasm. There are endogenous Ca2þ‐binding proteins (CaBPs) that provide buffer capacity as they bind Ca2þ with high affinity. Many CaBPs such as calmodulin, parvalbumin, calbindin‐D28K, and calretinin have helix–loop–helix (also called EF‐hand) motifs specific for Ca2þ binding in neurons (Baimbridge et al., 1992) and, therefore, play a neuroprotective role in CNS injuries and diseases (Goodman et al., 1993; Khachaturian, 1994). However, a decrease in expression of CaBPs in neurodegenerative processes may deprive the neurons from the protective actions of CaBPs against neuronal dysfunction and death.

2.5 Calpain Activation and Therapeutic Opportunity The complexity of the mechanisms leading to intraneuronal Ca2þ overload immediately suggests that it may be efficacious to block pharmacologically a common consequence of intraneuronal Ca2þ overload rather than focusing on one of the many Ca2þ overloading mechanisms (> Figure 15‐1). Because neurodegeneration with calpain activation is the ultimate consequence of intraneuronal Ca2þ overload (> Figure 15‐1), there exists an extended ‘‘window of opportunity’’ for use of therapeutic agents for calpain inhibition to prevent delayed neuronal death in CNS injuries and diseases.

3

Calpain Involvement in Neuronal Death in CNS Injuries

Major CNS injuries are ischemic brain injury, SCI, and TBI. Various studies suggest that calpain activation is an important part in progressive neurodegeneration in these CNS injuries. Therefore, calpain inhibition is considered to be an attractive therapeutic strategy in CNS injuries.

3.1 Ischemic Brain Injury Ischemic brain injury is an acute neurological disorder caused by blockage of cerebral blood supply, which leads to reduced availability of glucose and oxygen in the affected area. This is a condition of cellular energy crisis. Transient ischemic attacks exhibiting short duration of neurological dysfunction usually not exceeding 10–15 min may be a warning sign of impending stroke. Shortage of cellular energy interrupts the activity of ion pumps and thereby increases intracellular Ca2þ and extracellular Kþ concentrations, within 1–2 min after the onset of ischemia. Early ischemic damage is reversible to some extent. However, continuation of the ischemic condition can rapidly cause extensive irreversible damage due to onset of the ischemic cascade in the ischemic core. Ischemic cascade also affects areas around the infarct, leading to the formation of a so‐called ischemic penumbra. Prevention of neuronal death in the ischemic penumbra is the main target of pharmacological interventions.

3.1.1 Necrotic and Apoptotic Neuronal Death Ischemic brain injury triggers a number of cellular and molecular mechanisms for the mediation of neuronal death. Excitotoxicity, increase in intracellular free Ca2þ levels, cytokine‐mediated inflammatory responses, and scarcity of survival factors following cerebral ischemia can lead to neuronal death, which can be categorized into necrosis and apoptosis. Necrosis is associated with depletion of cellular energy, loss of membrane integrity, and random fragmentation of genomic DNA; while apoptosis is an ATP‐dependent and gene‐regulated cell death process characterized by appearance of apoptotic bodies, shrunken cytoplasm, condensed chromatin, and internucleosomal fragmentation of genomic DNA. Necrotic neuronal

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death predominantly occurs at the initial site of ischemic injury. Apoptotic neuronal death typically occurs at the periphery of the ischemic zone, also known as ischemic penumbra. Administration of neuroprotective agents in time after ischemic insult may rescue the neurons that succumb to apoptosis in the ischemic penumbra.

3.1.2 Calpain Activation in Neuronal Death and Calpain Inhibition A sustained increase in intracellular free Ca2þ level implied that Ca2þ‐dependent enzymes including calpain could be activated following ischemic brain injury to contribute to neurodegeneration (Liebetrau et al., 1999; Lipton, 1999; Neumar et al., 2001). Degradation of a‐fodrin and microtubule‐associated protein 2 (MAP2), two well‐known calpain substrates, preceded neurodegeneration in ischemic brain injury indicating that calpain was indeed involved in the early events triggering the cascades leading to neuronal death (Blomgren et al., 1995). Quantitative data also indicated that calpain‐mediated proteolysis occurred very early after the ischemic insult, correlated with the earliest changes in cell hypotrophy, and preceded or occurred in tandem with evidence of significant loss of neurons (Bartus et al., 1995a). Postischemic administration of the calpain inhibitor MDL‐28170 could ameliorate brain damage in animal models of ischemia (Li et al., 1998; Markgraf et al., 1998), confirming a prominent role for calpain in the pathophysiology of ischemic brain injury. Recent studies again suggest that calpain inhibition can prevent neuronal death after ischemic brain injury in mice (Jiang et al., 2005; Hou et al., 2006). It has been reported that the proteases of the calpain and caspase families act synergistically in neuronal apoptosis following ischemic brain injury (Rami et al., 2000; Yamashima, 2000; Blomgren et al., 2001; Zhang et al., 2002).

3.2 Spinal Cord Injury Spinal cord injury (SCI) occurs when mechanical trauma transfers energy through the complicated membrane system and fluid to the soft tissue of the spinal cord. Primary injury disrupts cell membrane, destroys axon–myelin structure in longitudinal tracts, and damages microvessels, initiating a devastating secondary injury process so as to release various detrimental factors that induce neuronal death. Multiple cellular and molecular mechanisms of the secondary injury process work though complex cascades over time to spread neurodegeneration beyond the site of primary injury.

3.2.1 Ca2þ Accumulation, Calpain Activation, and Neuronal Death A primary injury to the spinal cord increases intracellular free Ca2þ concentration in the SCI lesion (Happel et al., 1981). The increase in intracellular free Ca2þ concentration is due to a decrease in extracellular free Ca2þ concentration following SCI (Stokes et al., 1983; Xu et al., 1990). Increases in intracellular free Ca2þ levels (Happel et al., 1981; Du et al., 1999), reactive oxygen species (ROS) (Hall and Braughler, 1989; Braughler and Hall, 1992; Azbill et al., 1997; Yamamoto et al., 1998; Liu et al., 1999; Lewen et al., 2000), and lipid inflammatory mediators (Hsu et al., 1985; Xu et al., 1990) can stimulate calpain activity and mediate neuronal apoptosis in SCI. Calpain activation following SCI may have detrimental consequences (Banik et al., 1999) since many cytoskeletal and membrane proteins in neurons are well‐known calpain substrates, and they are readily degraded by calpain (Guttman and Johnson, 1999). We previously reported a requirement of de novo synthesis of proteins or proteases for cell death in the rat SCI lesion (Ray et al., 2001b), and involvement of calpain activity in the degradation of a 68‐kDa neurofilament protein (NFP) leading to internucleosomal DNA fragmentation in the SCI lesion and ischemic penumbra formation (Ray et al., 2000b). Thus, damage to the spinal cord did not remain confined to the site of injury or lesion, but extended (with the 68‐kDa NFP degradation and DNA fragmentation) over time to the rostral and caudal regions.


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3.2.2 Calpain Inhibition for Neuroprotection Calpain activation and neurodegeneration in SCI warranted examination of the therapeutic efficacy of calpain inhibitors in neuroprotection. Calpastatin (110 kDa), the endogenous protein inhibitor of calpain, is too large to be cell permeable. Also, with an increased calpain/calpastatin ratio, calpain degrades calpastatin as a suicide substrate in vitro (Mellgren et al., 1986; Nakamura et al., 1989), in cultured cells (Nagao et al., 1994), and even in vivo (Saido et al., 1997; Blomgren et al., 1999). Therefore, the therapeutic use of calpastatin for the inhibition of calpain in SCI is not feasible. However, the use of calpain‐specific inhibitors has been beneficial in several pathophysiological conditions. For example, E‐64‐d was the first reported cell‐permeable calpain inhibitor (Tamai et al., 1986) that reduced muscle degeneration in animal models of muscular dystrophy (Komatsu et al., 1986; Tamai et al., 1987), calpeptin and MDL‐28170 improved posthypoxic recovery of cells (Arlinghaus et al., 1991), and AK295 protected neurons in brain ischemia (Bartus et al., 1994). We have recently described in detail the involvement of calpain in the pathophysiology of SCI and the potential of calpain inhibitors for neuroprotection in the lesion and penumbra following SCI (Ray et al., 2003). Studies from other laboratories also suggest that inhibition of calpain activity provides beneficial effects in SCI animals (Du et al., 1999; Schumacher et al., 2000). Recent studies also report that calpain inhibitors prevent neuronal death and improve neurological function following SCI in rats (Arataki et al., 2005; Hung et al., 2005). These findings in animal models of SCI strongly support the therapeutic use of calpain inhibitors in providing functional neuroprotection following SCI in humans.

3.3 Traumatic Brain Injury Traumatic brain injury (TBI) is triggered by an external mechanical impact to the head. After this primary impact to the head, a secondary injury process is developed over hours or days so as to release neurodestructive compounds. Multiple mediators of the secondary injury process ultimately cause activation of cysteine proteases that cleave key cellular substrates, leading to neuronal death.

3.3.1 Changes in Ca2þ Homeostasis The neuropathology of TBI is complex. Increases in intracellular free Ca2þ levels have been reported in brain regions after TBI (Shapira et al., 1989; Fineman et al., 1993). A number of other studies indicate that elevated intracellular free Ca2þ plays an important role in the initiation of pathophysiological pathways leading to neurodegeneration after CNS trauma (Tymianski and Tator, 1996; McIntosh et al., 1997). It has been proposed that activated ion channels following TBI may contribute to prolonged changes in Ca2þ homeostasis (Gennarelli et al., 1998). Therefore, Ca2þ channel blockers have been examined as possible neuroprotective agents to reduce excessive accumulation of intracellular free Ca2þ levels in experimental TBI. Administration of voltage‐sensitive Ca2þ channel blockers provided little success in treating brain injury in humans (Baethmann and Jansen, 1986; Robinson and Teasdale 1990). Dihydropyridine Ca2þ channel blockers like nicardipine and nimodipine have been evaluated in clinical trials in TBI. Early trials reported beneficial effects of nimodipine in severe head injury in humans (Kostron et al., 1984). In contrast, more recent trials with both nicardipine and nimodipine did not show any clinical benefits in patients with TBI (Compton et al., 1990; Teasdale et al., 1992). Thus, the therapeutic efficacy of Ca2þ channel blockers in treating TBI remains controversial.

3.3.2 Calpain Activation and Calpain Inhibition The important fact is that an increase in intracellular free Ca 2þ levels following TBI activates the Ca2þ‐dependent-cysteine protease calpain, leading to neurodegeneration (Kampfl et al., 1996; Ray et al.,

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2002). The Ca2þ‐independent cysteine proteases, especially caspase‐3, can also play a role in mediating cell death following TBI. Although activation of caspase‐3 via the extrinsic and intrinsic apoptotic pathways after moderate TBI has been documented (Beer et al., 2000; Keane et al., 2001), experimental studies strongly suggest that calpain is more important than caspase‐3 for the mediation of cell death after TBI (Kampfl et al., 1997; Pike et al., 1998; Pike et al., 2001; Ray et al., 2002). Calpain activation may occur upstream of caspase‐3 for the mediation of apoptosis as suggested by both in vitro (Waterhouse et al., 1998) and in vivo (Ray et al., 2001a) studies. The increase in intracellular Ca2þ after TBI certainly activates Ca2þ‐ dependent proteases including calpain, which mediates cytoskeletal protein degradation and neurodegeneration in humans (McCracken et al., 1999; Huang and Wang, 2001) and rodents (Kampfl et al., 1996, 1997; Saatman et al., 1996a; Pike et al., 2001; Ray et al., 2002). Recently, differential proteome analysis indicated the involvement of calpain and caspase‐3 in the degradation of various substrates following TBI (Liu et al., 2006). Some studies used calpain inhibitors such as calpeptin and MDL‐28170 to demonstrate the involvement of calpain in apoptotic death of rat glial (Ray et al., 1999) and neuronal (Ray et al., 2000) cells, respectively. Other calpain inhibitors such as calpain inhibitor II, AK295, and SJA6017 in rodent models attenuated the loss of cytoskeletal proteins after cortical impact brain injury (Posmantur et al., 1997) and improved functional outcome after fluid‐percussion brain injury (Saatman, et al., 1996b) and diffuse brain injury (Kupina et al., 2001). Calpain inhibitors are capable of providing neuroprotection both in vitro and in vivo models suggesting that calpain inhibition can be an important therapeutic strategy in TBI.

4

Calpain Involvement in Neuronal Death in CNS Diseases

Calpain activation in the mediation of neuronal death has also been demonstrated in a number of CNS diseases. There is an increasing effort in using calpain inhibitors for preventing neuronal death in cell culture and animal models of CNS diseases. Recent results suggest that calpain can be a promising target for the prevention of neurodegeneration in many major CNS diseases.

4.1 Alzheimer’s Disease The most common neurodegenerative disease worldwide is Alzheimer’s Disease (AD), which is characterized by loss of hippocampal neurons and synapses resulting in cognitive impairment, loss of memory, lack of language skills, and reduced reasoning leading to dementia and finally death (Selkoe, 2004).

4.1.1 Role of Calpain in Neurodegeneration A role for calpain in the pathogenesis of AD has been suspected more than a decade ago (Nilsson et al., 1990). Calpain may be involved in the formation of b‐amyloid peptides and pathogenesis of AD (Siman et al., 1990). In another study, an intense immunoreactivity of calpain in the dystrophic neurites of senile plaques in brains from patients with AD suggested that calpain could be involved in the pathogenesis of AD (Shimohama et al., 1991). Furthermore, double staining experiments revealing calpain immunoreactivity in senile plaques and in neurons undergoing neurofibrillary degeneration in AD strongly suggested that activation of calpain could be an important factor in the abnormal proteolysis underlying the accumulation of plaques and tangles in AD (Iwamoto et al., 1991). As an index of change in the in vivo activity of calpain in AD, the ratio of the 76‐kDa active fragment of calpain to its 80‐kDa precursor was measured by immunoassay in selected brain regions from individuals with AD and normal controls (Saito et al., 1993). This activation ratio was elevated three-fold in the prefrontal cortex from patients with AD and it was also significantly elevated, but to a lesser degree, in brain regions where AD pathology was milder, indicating that calpain activation was a potential molecular basis for neuronal degeneration (Saito et al., 1993). Further studies using active site‐directed antibodies identified calpain as an early‐appearing and pervasive component of neurofibrillary pathology in AD (Grynspan et al., 1997). Therefore, pharmacological modulation of calpain activity should be considered as a potential therapeutic strategy for treatment of AD.


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4.1.2 Calpain Inhibition as a Therapeutic Strategy Treatment of primary cortical neurons with the amyloid b‐peptide Ab(1‐42) activates calpain for the conversion of p35 to p25, which is also accumulated in the brains of patients with AD, suggesting that calpain may be involved in the pathogenesis of AD (Lee et al., 2000). However, a subsequent study shows that p35 is rapidly cleaved to p25 in rat and human brains within a short postmortem delay, and immunoblot analysis of brains prepared from patients with AD or age‐matched control individuals with a short postmortem delay reveals no specific increase in the levels of p25 in the brains of patients with AD (Taniguchi et al., 2001). In contrast, levels of active form of calpain were increased in brains of patients with AD compared with those in controls, suggesting that the conversion of p35 to p25 is a postmortem degradation event and may not be upregulated in AD brains (Taniguchi et al., 2001). Various studies show how calpains, acting directly or indirectly through other proteolytic pathways and cellular signaling cascades, may promote b‐amyloidogenesis, neurofibrillary pathology, and mediate neurodegeneration in AD (Nixon, 2000). Confocal immunofluorescence study with antibodies specifically recognizing the active form of calpain provided evidence of the important role of the calpain proteolytic system in the pathogenesis of AD (Adamec et al., 2002). Current understanding of the molecular mechanisms in AD strongly suggests calpain activation in neurodegeneration and calpain inhibition as a possible approach for the treatment of AD (Higuchi et al., 2005). Investigations have been conducted to explore calpain inhibitors as potential therapeutic agents for the treatment of AD (Di Rosa et al., 2002).

4.2 Amyotrophic Lateral Sclerosis Amyotrophic lateral sclerosis (ALS), also called Lou Gehrig’s disease, is a progressive neurodegenerative disease neuropathologically characterized by selective loss of motoneurons mainly in the spinal cord and in some cases in the brainstem and brain (Rowland, 1995). Most cases of ALS are sporadic and about 10% of all cases of ALS are familial. Both forms of ALS are indistinguishable in terms of clinical and neuropathological features (Rowland, 1995).

4.2.1 Mechanism of Motoneuron Death The exact mechanism contributing to the pathogenesis of sporadic ALS is unknown, but several observations suggest that glutamate toxicity may participate in selective motoneuron degeneration (Rothstein et al., 1992; Shaw and Ince, 1997). Glutamate‐mediated overactivation of NMDA receptor opens the divalent ion channels to allow a substantial influx of Ca2þ into the cell (Mayer et al., 1984). Overactivation of a subtype of AMPAR/KAR by glutamate may also increase the permeability of Ca2þ (Iino et al., 1990). Spinal cord motoneurons indeed possess Ca2þ‐permeable AMPAR/KAR (Bar‐Peled et al., 1999). Another type of glutamate receptor is the metabotropic receptor that, unlike ionotropic receptors, is associated with the activation of an intracellular second messenger. Binding of glutamate to this type of receptor activates phospholipase C (Wei et al., 1982), which may induce the synthesis of IP3 for releasing Ca2þ from the intracellular stores (Sugiyama et al., 1987).

4.2.2 Calpain Activation in Motoneuron Death and Calpain Inhibition Alterations in intracellular free Ca2þ concentrations may be crucial to the diverse pathogenic mechanisms potentially responsible for ALS (Krieger et al., 1996). Also, studies provide evidence that oxidative damage may play a role in the pathogenesis of neuronal degeneration in both sporadic and familial ALS (Ferrante et al., 1997). Both oxidative stress and Ca2þ influx upregulate calpain and induce apoptotic death of

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neurons (Ray et al., 2000; See and Loeffler, 2001). The mRNA levels of calpain 1 and calpain 2 were significantly increased in patients with ALS as compared with controls, suggesting that calpain could likely be playing a regulatory role in the process of myofibrillar degradation in ALS (Ueyama et al., 1998). A previous study indicated that PD150606, an inhibitor of calpain (Wang et al., 1996), prevents selective motoneuron death via inhibition of Ca2þ influx (Van den Bosch et al., 2002). Recently, we reported that calpain played a role in motoneuron death and calpain inhibitor provided functional protection to motoneurons (Das et al., 2005), implying that calpain inhibition could be a potential therapeutic strategy for functional protection of motoneurons in ALS.

4.3 Epileptic Seizures Epilepsy, which may occur at any age, is a group of neurological disorders characterized by chronic and recurrent disturbances in neurological functions due to abnormalities in electrophysiological activity in the brain, leading to neuronal death and dysfunction (Leppik, 1986; Henshall and Simon, 2005). Each episode of neurodysfunction is known as seizure. Studies indicate that epileptic seizures reflect cerebral disturbances due to dysfunction of diverse fundamental neurochemical mechanisms that involve alterations in excitatory and inhibitory functions, resulting in hyperexcitability hypersynchronization, or both. Neurological manifestations of epileptic seizures vary from a brief lack of attention to a prolonged loss of consciousness with convulsive motor activity.

4.3.1 Excessive Intracellular Free Ca2þ and Calpain in Pathogenesis Excessive influx of Ca2þ due to repeated depolarization and prolonged stimulation of NMDAR is proposed to be the ultimate cause of neuropathology in nerve agent‐induced seizures (McDonough and Shih, 1997). The colocalization of neuronal damage with demarcated reductions in MAP2 in rat models of seizure represented the first demonstration of a sensitive marker in seizure‐induced brain damage (Ballough et al., 1995), and indicated the involvement of proteases in this process. A subsequent study identified an association of increased calpain activation with neuropathology following seizure onset in adult rat brains (Bi et al., 1996). Other studies show that hippocampal dentate granule neurons and CA1 pyramidal neurons, which express the Ca2þ‐buffering protein calbindin D28K, may be less vulnerable to damage following kainate‐induced seizure in mice (Gary et al., 2000), and support the notion that Ca2þ‐dependent proteases such as calpain may be involved in the pathogenesis of seizure‐induced injury (Vanderklish and Bahr, 2000). Further studies suggest that calpain participates in the molecular mechanisms of seizure‐ induced cell death (Kondratyev and Gale, 2004).

4.3.2 Oxidative Stress Leading to Calpain Activation Oxidative stress is emerging as a mechanism that may play an important role in the initiation of epileptic seizures and neuronal death (Patel, 2002). Mice heterozygous for oxygen‐regulated protein‐150 kDa (ORP150), a molecular chaperone present in ER, have been used in a recent study showing that exposure to excitatory stimuli caused hippocampal neurons to display exaggerated elevation of cytosolic free Ca2þ levels and activation of calpain accompanied by increased vulnerability to glutamate‐induced cell death in vitro and decreased survival to kainate‐induced seizures in vivo (Kitao et al., 2001). This report therefore provides a mechanism that ORP150 regulates cytosolic free Ca2þ levels and the activation of the calpain pathway causing cell death in hippocampal neurons following induction of seizures. However, no recent experimental results are yet produced showing the efficacy of newly discovered calpain inhibitors in seizure‐ induced neuronal death.


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4.4 Huntington’s Disease Huntington’s disease (HD) is an inherited autosomal dominant neurodegenerative disease, which is clinically characterized by uncontrollable movements and restlessness (termed ‘‘chorea’’), irritability, and depression (Margolis and Ross, 2003). The age of HD onset is between 20 and 50 years and death usually occurs within 15–20 years from the beginning of clinical symptoms (Mendez, 2004). There has been exciting progress in recent years in unraveling several potential mechanisms of pathogenesis in HD (Petersen et al., 1999), which is neuropathologically characterized by selective death of cortical neurons and development of astrogliosis in the striatum (MacMillan and Quarrell, 1996; Vonsattel and DiFiglia, 1998). HD affects all races although it has the highest prevalence in Europe and North America (Harper, 1992).

4.4.1 Mechanism of Pathogenesis Huntington’s disease is a CAG repeat or polyglutamine disease as it is caused by an expansion of the trinucleotide CAG (codon for glutamine) in the HD gene resulting in a mutant form of a protein called huntingtin, whose normal function is not known (Gusella et al., 1983; Huntington’s Disease Collaborative Research Group, 1993; Mangiarini et al., 1996). The pathogenesis of HD is presumed to be due to a toxic gain‐in‐function of the mutant huntingtin (Petersen et al., 1999) with the involvement of excitotoxicity (Coyle and Schwarcz, 1976; Beal et al., 1986), oxidative stress (Coyle and Puttfarcken, 1993), and impaired energy metabolism (Beal, 1992; Jenkins et al., 1993; Dragunow et al., 1995; Portera‐Cailliau et al., 1995; Thomas et al., 1995; Gu et al., 1996).

4.4.2 Excitotoxicity, Ca2þ Influx, and Calpain Activation Glutamate‐mediated excitotoxicity plays a crucial role in the pathogenesis of HD. Ionotropic glutamate receptors such as NMDAR (which are sensitive to NMDA as well as glutamate and quinolinic acid), AMPAR (which are sensitive to AMPA and kainic acid), and kainic acid/quisqualic acid receptors (which are sensitive to kainic acid and quisqualic acid) control ion channels and contribute to neurodegeneration (Lipton and Rosenberg, 1994). Upon excitotoxic activation, the NMDAR channels allow Ca2þ influx. Also, mGluR are coupled to G‐proteins and potentiate NMDAR‐mediated Ca2þ influx via protein kinase C activation, promoting the pathways toward neuronal death (Schoepp and Sacaan, 1994). An increase in Ca2þ influx in neurons can activate phospholipase A2 (PLA2)‐ and Ca2þ‐dependent proteases including calpain. Activation of PLA2 increases production of arachidonic acid, and the subsequent metabolism of arachidonic acid leads to the formation of free radicals, and thus contributing to neurotoxicity. In recent years, accumulating direct or indirect evidence strongly suggests that activation of calpain can contribute to neurodegeneration in HD (Kim et al., 2001; Gafni and Ellerby, 2002; Goffredo et al., 2002). A recent study showed that calpain‐mediated fragmentation of mutant huntingtin played a role in the pathogenesis of HD and calpain inhibition reduced the toxic accumulation of mutant huntingtin fragments in the striatum (Gafni et al., 2004).

4.5 Parkinson’s Disease Parkinson’s disease (PD) or parkinsonism is a major movement disorder disease. It refers to a clinical condition with symptoms that mainly include tremor, rigidity, bradykinesia, and imbalance (Lang and Lozano, 1998). PD is neuropathologically characterized by the degeneration of dopaminergic neurons in the substantia nigra (Hirsch et al., 1989; Forno, 1996), a particular area of the brain critical for controlling movement.

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4.5.1 a‐Synuclein and Oxidative Stress Lewy in 1914 first described the presence of neuronal inclusion bodies (now known as Lewy bodies) in the substantia nigra of PD patients (Lewy, 1914). a‐Synuclein is a major component of the Lewy bodies (Spillantini et al., 1997; Wakabayashi et al., 1997). A recent study suggests how a‐synuclein aggregation may occur in a vicious cycle causing neurotoxicity and eventually neuronal death (Liu et al., 2005). Some cases of PD include other pathologies with the involvement of a certain system of nerve cells scattered throughout the brain and spinal cord (Jellinger, 1987; Paulson and Aotsuka, 1993). There is a concern that oxidative stress involving H2O2 and free radicals may contribute to the pathogenesis of PD (Olanow, 1990). Dopaminergic neurons of substantia nigra are particularly vulnerable to oxidative stress. Other plausible mechanisms of neuronal cell death in PD are also being explored.

4.5.2 Calpain Activation and Calpain as a Target for Therapy Activation of NMDAR in PD may induce excitotoxicity that causes accumulation of intracellular free Ca2þ leading to activation of calpain. The NMDAR antagonists reduce the depletion of dopaminergic neurons in substantia nigra and prevent the development of parkinsonism in animal models (Turski et al., 1991), suggesting a role for excitotoxicity in the pathogenesis of PD. The Ca2þ‐buffering protein calbindin plays a neuroprotective role by regulating intracellular Ca2þ concentrations (Mattson et al., 1991). Dopaminergic neurons expressing the Ca2þ‐buffering protein calbindin are selectively preserved in the substantia nigra of patients with PD (Hirsch et al., 1992), suggesting that the neurons which degenerate in PD are particularly sensitive to a rise in intracellular free Ca2þ concentrations and Ca2þ‐dependent proteases such as calpain leading to neuronal cell death in PD. An immunohistochemical analysis revealed an increase in calpain expression in the mesencephalon of patients with PD and suggested the involvement of calpain in neuronal death in PD (Mouatt‐Prigent et al., 1996). This notion is subsequently strengthened with an observation that calpain overexpression in PD is not compensated for by a concomitant increase in the expression of calpastatin (Mouatt‐Prigent et al., 2000), the endogenous calpain inhibitor. Upregulation of calpain indeed can play a role in neuronal death in brain and spinal cord of mouse models of PD (Ray et al., 2000; Chera et al., 2002; Chera et al., 2004). A recent study showed a prominent role for calpain in neuronal apoptosis in experimental models of PD and also in neuroprotection after calpain inhibition (Alvira et al., 2006). Inhibition of calpain as a therapeutic strategy needs to be further confirmed in animal models of PD.

4.6 Multiple Sclerosis Multiple sclerosis (MS) mostly affects women. It is an autoimmune demyelinating disease of the CNS (Ffrench‐Constant, 1994; Sorensen and Ransohoff, 1998; Antel, 1999) in which the myelin and myelin‐ producing oligodendroglial cells become the target of an inflammatory response, resulting in depletion of the white matter (Williams et al., 1994; Raine, 1997).

4.6.1 Multiple Mechanisms of Pathogenesis Some studies support the importance of genetics in determining the risk of developing MS (Ebers et al., 1995; Oksenberg et al., 1996; Sadovnick et al., 1996). No abnormal or mutant genes have yet been conclusively linked to MS, but polymorphic variants of some genes that determine specific immune responses have been implicated in the pathogenesis of MS (Seboun et al., 1989; Utz et al., 1993; Sawcer et al., 1996; Yaouanq et al., 1997). The neuropathological hallmark of MS remains to be the white matter plaque, a defined patch of demyelination, which is widely believed to be due to the destruction of myelin proteins (Martin and McFarland, 1995) in response to an autoimmune inflammatory reaction mediated by


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T cells following their infiltration into the CNS (de Rosbo and Ben‐Nun, 1998). The strong support for considering MS as an autoimmune disease comes from studies with its animal model, experimental allergic encephalomyelitis (EAE) (Moore et al., 1984; Zamvil and Steinman, 1990; Linington et al., 1993). Studies on EAE reveal that TH cells (CD4þ, MHC II restricted) recognize myelin antigens presented by mononuclear phagocytes (macrophages, microglia, and dendritic cells), triggering an autoimmune inflammatory cascade that culminates in the destruction of myelin proteins.

4.6.2 Potential Role of Calpain in Pathogenesis Degradation of various myelin proteins in MS implies the involvement of proteases in the pathogenesis of this disease. The possibility of the involvement of a Ca2þ‐dependent neutral protease in the degradation of myelin proteins in MS was postulated more than 20 years ago (Banik et al., 1985; de Rosbo and Bernard, 1989). Later an in vitro study in our laboratory indicated that activated human lymphoid or monocytic cells secrete calpain to cleave purified myelin basic protein (MBP) and myelin (Deshpande et al., 1995), implying that calpain‐cleaved MBP products could include fragments with antigenic epitopes that play an important role in the initiation and perpetuation of immune‐mediated demyelination. Support for the involvement of calpain and other proteolytic enzymes in the process of demyelination came from different laboratories (Smith, 1979; Smith et al., 1998). Our laboratory also reported the increased calpain expression and activity in demyelinating disease (Shields and Banik, 1998; Shields et al., 1998; Schaecher et al., 2001) and an interconnection among calpain upregulation, extent of myelin loss, and severity of the disease in EAE rats (Guyton et al., 2005b) and postmortem tissue from patients with MS (Shields et al., 1999). Current investigations indicate that MS is not only a demyelinating disease but it also has a neurodegenerative component as axons and neurons are affected (Ferguson et al., 1997; Trapp et al., 1998; Peterson et al., 2001). Although the mechanisms by which axons and neurons are damaged are not clearly known, elevation of intracellular free Ca2þ and calpain activation have strongly been implicated in the neurodegenerative process in EAE animals and MS patients (Guyton et al., 2005a). Other investigators demonstrated an association of calpain activity with age‐dependent myelin degeneration and proteolysis of oligodendrocyte proteins in rhesus monkeys that developed significant cognitive impairment (Sloane et al., 2003). Presently, there is a lack of effective therapy for MS because the underlying cause of the disease and many other critical features of the etiology and pathogenesis remain mostly unidentified. However, increasing evidence of calpain involvement in animal models of demyelinating disease suggests that calpain inhibitors may be tested as potential therapeutic agents for the treatment of demyelinating diseases.

5

Conclusion and Future Directions

Over the years, many investigators identified calpain activation in the pathophysiology of a wide variety of CNS injuries and diseases. The involvement of calpain in CNS injuries and diseases has already created great interest for the development of highly potent, selective, and cell‐permeable calpain inhibitors for therapeutic applications. It should be emphasized that there are difficulties in developing water‐soluble small molecule calpain inhibitors with appropriate pharmacological properties. Extensive research continues to identify therapeutically useful calpain inhibitors for the prevention of neurodegeneration in CNS injuries and diseases.

Acknowledgments This work was supported in part by the R01 grants (CA‐91460, NS‐31622, NS‐38146, NS‐41088, NS‐45967, and NS‐57811) from the National Institutes of Health and also a Spinal Cord Injury Research Foundation grant (SCIRF‐0803 and SC9RF‐1205) from the state of South Carolina.

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Secretase Processing of Amyloid Precursor Protein (APP) and Neurodegeneration

N. Marks . M. J. Berg

1 1.1 1.2 1.3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 470 The Amyloid Hypothesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 471 APP Heterogeneity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 473 Regulated Intramembrane Proteolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475

2 2.1 2.2 2.3 2.4 2.5

a-Secretases (K16L17 Cleavage) and the ADAM Gene Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 Domain Structure and Therapeutic Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 477 Constitutive and Regulated APP Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 478 Alternative a-Secretases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 478 Therapeutics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479

3 3.1 3.1.1 3.2 3.3

b-Secretases (BACE-1 and -2) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479 Domain Organization and Implications for Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 480 Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 480 Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 481 Pharmacology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 481

4 4.1 4.2 4.3 4.4 4.4.1 4.5 4.6

g-Secretase Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 483 Clarifying g-SC as a Multimeric Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 483 Stepwise Assembly and Maturation of Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 483 Isolation of g-SC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 485 Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 487 AICD and Cell Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 489 g-SC Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 490 BRI Amyloidoses and lnhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 492

5 5.1 5.2 5.3 5.4

Obligatory Components of the g-SC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 493 Presenilins (PS1, PS2) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 493 Nicastrin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 495 Aph-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 495 Pen-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 496

6

Alternative Proteases as Secretases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 496

7

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 497

#


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Secretase processing of amyloid precursor protein (APP) and neurodegeneration

Abstract: Secretases releasing fibril-forming fragments from Amyloid Precursor Protein (APP) that accumulate in extracellular deposits in Alzheimer’s disease (AD) and parallel cognitive dysfunction, are described according to their historical discovery. These focus on metalloproteases that link to normal turnover (α-, ADAMs), a new family of Asp proteases (β-, BACE) for release of an intermediate for final cleavage by γ- to release amyloid β-peptide (Aβx). While α- and β-secretases are available in purified forms for rational synthesis of inhibitors, γ- exists only as a partially purified tetrameric complex (γ-SC) containing Presenilin, Nicastrin, Aph-1 and Pen-2 as obligatory components. Wide specificity of secretases precludes design of clinical agents to specifically target APP fragmentation. γ-SC cleavage of membranebound proteins points to important roles for Regulated Intramembrane Proteolysis (RIP) for both protein turnover and release of modulators of diverse signaling pathways. The review considers controversies regarding APP processing in sporadic AD (SAD), and the role of mutations for APP or Presenilin to exacerbate fibrillogenesis in familial AD (FAD). Progress in characterization of secretases provides a new basis for novel AD therapies and insights in general mechanisms of cellular protein turnover. List of Abbreviations: ADAM, A Disintegrin And Metalloproteases; ACDL, acidic dileucine; APP, amyloid precursor protein; Aph, anterior pharynx defective phenotype; APLP, APP-like proteins; AICD, APP intracellular domain; AD, Alzheimer’s disease; Aβx, Amyloid peptide, 4 kDa with variable length 39-43 residues; BACE, β-APP-Converting Enzyme; CD, cerebrovascular deposits; CAA, cerebral amyloid angiopathy; DS, Down’s syndrome; FAD, Familial Alzheimer’s disease; γ–SC, γ–secretase complex; IDE, insulin degrading enzyme; ICD, intracellular domain; KO, knock-out; NCT, Nicastrin; NSAIDS,, non-steroidal anti-inflammatory drugs; NICD, Notch intracellular domain; pen-2, presenilin enhancer 2; PS, Presenilin; SAD, Sporadic Alzheimer’s disease; SP, senile plaque; RIP, Regulated Intramembrane Proteolysis; SP, signal peptide; TIMP, Tissue Inhibitors of Metalloproteases; Tg, transgenic; TM, transmembrane domain; TNF-α, tumor necrosis factor α; TACE, tumor necrosis factor α converting enzyme

1

Introduction

Secretases are named since they convert amyloid precursor protein (APP) for secretion as metabolites, which are of interest in Alzheimer’s disease (AD) and cell signaling pathways (> Table 16-1). Notably they are responsible for the release of a 4-kDa fragment (amyloid peptide, Abx), which is a major component of senile plaque (SP) and cerebrovascular deposits (CD). Abx results from sequential processing by b- and then g-secretase while a third enzyme, a-secretase, destroys the fibril-forming consensus sequence and is nonamyloidogenic (> Figure 16-1). Interest in secretases grew after the isolation of the 4-kDa fragment of the then unknown APP in 1984 by Glenner and Wong (Glenner and Wong, 1984a, b), or roughly 80 years after the pathological description of abnormal argentophilic deposits by Alzheimer in 1907 (Alzheimer, 1907). Remarkably Glenner and Wong succeeded in identifying the 4-kDa polypeptide in the insoluble buffy layer by using simple hypotonic extracts of leptomeningeal membranes from patients with Alzheimer’s disease and Down’s syndrome (AD/ DS) or SP-enriched tissue. This seminal finding rapidly led to the following developments of importance to the present theme: Abx as the major component of SP (25% of dry weight) Cloning and characterization of APP Association with missense APP mutations and another family of novel proteins termed presenilins (PS) in familial AD (FAD), which represents in toto only 5% of all cases with dementia FAD proteins that were used for creating transgenic (Tg) models or for transfection Isolation of b-secretases (BACE-1/2) as a novel family of Asp proteases New functions for metalloproteases of the ADAM (A Disintegrin And Metalloprotease) family as putative a-secretases Processing of a cell-associated CTF by multimeric complexes (g-SC)


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. Table 16-1 Terminology of secretases Site (secretase) a

b

g-Secretase complex (g-SC)c

Terminology/comments TACE (ADAM-17) (Black et al., 1997; Moss et al., 1997)KUZ (ADAM-10) (Lammich et al., 1999), other ADAM family members BACE-1/2 (Sinha et al., 1999; Vassar et al., 1999; Abbenante et al., 2000) Alternative nomenclature: Asp-1/2 (Hussain et al., 1999) Memapsin-1/2 (Hussain et al., 1999; Lin et al., 2000) Presenilin (PS-1 or -2) and NTF/CTF heterodimer (> Sect. 4.2 and 5.1) Nicastrin (NCT; Caenorhabditis elegans aph-2) (Goutte et al., 2000; Yu et al., 2000; Morais et al., 2003; Shirotani et al., 2003; Niimura et al., 2005) aph-1 and isoforms (Goutte et al., 2002), anterior pharynx defective phenotype (Shirotani et al., 2004) pen-2, presenilin enhancer-2 (Francis et al., 2002; Luo et al., 2003; Takasugi et al., 2003)

Representative activators/inhibitors Hydroxamatesa, chelating reagents, phorbol esters (Buxbaum et al., 1993), APMA KTEEISEVNLstatinVAEFb, OMOO-3/4 (Hom et al., 2004; Turner, III et al., 2005)

Hydroxyethylene dipeptide isotere L685,458 (Shearman et al., 2000)Miscellaneous: DAPT (Dovey et al., 2001), fenchylamine, sulfonamides (Rishton et al., 2000), difluoroketone peptidomimetics (Moore et al., 2000), NSAIDs (Marambaud et al., 1998; Wolfe et al., 1999; Skovronsky et al., 2000b; Beher et al., 2004), benzodiazepines (Churcher et al., 2003; Owens et al., 2003), etc.

a

IC3, others include batimistat, marimastat (see text) GI-120471, SE-205, TAPI, KD-1X-73-4 (Newton and Decicco, 1999; Racchi et al., 1999; Parvathy et al., 2000) b Employed for affinity purification (Sinha et al., 1999) c For stepwise assembly of a putative quaternary g-SC ( > Sect 4.2) Abbreviations: TACE, tumor necrosis factor-a-converting enzyme; ADAM, a disintegrin and metalloendopeptidase; KUZ, kuzbanian protease; BACE, b-amyloid-converting enzyme; a-1PDX, a1-antitrypsin; L685,458, (1S-benzyl-4R-(1-(1S-carbamoyl-2-phenylethylcarbamoyl)-1S-3-methyl-butylcarbamoyl)-2R-hydroxy-5-phenylpentyl) carbamic acid tert butyl ester, a transition-state aspartyl-like inhibitor; NSAIDs, nonsteroidal antiinflammatory drugs (for sample structures > Table 16-4 or commercial websites for inhibitors such as Calbiochem/EMD Biosciences: http://www.emdbiosciences.com). > Table 16-5 for properties and structural determinants of obligatory components in g-SC. Some key references are supplied; in other cases see text under headings for secretases. For a full listing of the ADAM and BACE families, consult MEROPS database (http://delphi.phys.univ-tours.fr/Prolysis/merops/merops.htm)

1.1 The Amyloid Hypothesis The amyloid hypothesis proposes that the incidence of abnormal lesions correlates with the histopathology and cognitive dysfunction. It is believed that deposits trigger neuronal cell death and the hallmark loss of synapses and appearance of intracellular tangles. Abx peptides in solution form protease-resistant fibrils/ aggregates, which in situ may provide a nidus for the later accretion of cellular debris and account for the heterogeneous composition of SP that includes a variety of proteases or their inhibitors (cystatin, antichymotrypsin), proteoglycans and proteolipids (syndecan, ApoE), glycation products (RAGE), and metals, all of which are thought to exacerbate fibrillogenesis (Marks and Berg, 1997). Paradoxically, cystatin c, an inhibitor of lysosomal cysteine proteases, binds Ab and in vitro, interferes with its oligomerization (Sastre et al., 2004). However, toxicity may be exerted by plaques and A42 oligomers, altering Ca2þ homeostasis (Demuro et al., 2005). SPs are characterized by a progressive change in morphology that includes invasive neurites, neuronal threads, Hirano-like bodies, and granulovacuolar lesions surrounded by astrocytes/microglia, which contribute

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. Figure 16-1 Sites of APP cleavage by secretases. Domain structure in the center is for the APP770 isoform. FAD mutations are italicized and named on the basis of the original kindreds (--->). *Murine residues that differ from hAPP. The sequence in bold (top) reflects the Ab numbering. Exploded sequence (bottom) indicates site for caspase cleavage (D739) and a consensus sequence for interaction with nuclear adaptors or endocytic internalization. Dashed boxes represent putative transmembrane domain TM. AICD, amyloid intracellular domain; SP, signal peptide; TM, transmembrane; KPI, Kunitz protease inhibitor; ♦, phosphorylation site; , glycosylation site

multiple components. Dystrophic neurites, for example, contain APP, tau, dense-laminated bodies, neurofilaments, GAP-43, PKC, ubiquitin, spectrin, and neurotransmitters. AD deposits are distinctive and differ from those seen in Pick’s, Gerstmann–Straüssler, Parkinson’s, and Lewy body diseases, which form from diverse proteins (> Table 16-2) with mature plaques, but not amorphous ones, having b-pleated structures (birefringence, staining by silver salts, Congo red, and thioflavin). Many of these occur in association with Abx or show autosomal dominant mutations, suggesting that similar therapeutic approaches may result in the reduction of abnormal 3D conformations (Chiti et al., 2002) (> Figure 16-2). These approaches include the following: Using protease inhibitors, RNAi silencing, or use of antisense probes Using b-disruptive agents or immunotherapy (anti-Abx) Promoting normal physiological turnover (a-secretase) Altering the targeting of enzyme or substrate Diet (Calorie Intake: Scyllo-inositol supplemented) (McLaurin et al., 2006).


16

. Table 16-2 Toxic consequences of amyloid or other aggregates Fibril-forming entity Ab40-42 Fragment Amyloid ABr/Dan Dementias (British and Danish kindreds with mutant forms of BRI2, a type-2 transmembrane protein, > Sect 4.5)

Tau

a-Synuclein

Gln Repeat Disorders (CAG)

Prion (PrP) S. cerevisiae prion (PHGGGWGO repeats) Cystatin c

Pathology SAD/FAD FAD linking with Chromosome 13 with severe CAA. Amyloid fragments ABr/ADan. Amyloid A subtypes with identical N-terminus EASNCFAIRHFENKFAVITLIC* Br: 23SRTVKKNIIEEN34 Dan: 23FNLFLNSQEKHY34 Divergent C-termini arising from a point mutation in Br, 10 nt duplication in Dan Tangles (3–4 tau repeats) Pick’s Disease (FTDP-17) Frontal-Temporal Dementia w/Parkinsonism associated w/Chr 17 and tau gene) SAD/FAD Parkinson’s Disease A53T, ?A30P Progressive Supranuclear Palsy (PSP) Parkin (Ubiquitin Ligase) HD – Huntingtin (38–180 repeats) SBMA – Androgen R (38–65 repeats) SCA 1 – Ataxin-1 2 – Ataxin-3 (55–64 repeats) 7 – Ataxin-7 (24–200 repeats) DRPLA – Atrophin-1 (48-88 repeats) (nuclear inclusions?) Jacob-Creutzfeldt Disease Plaques and Lewy Bodies Spongiform Encephalopathies Hereditary Cerebral Hemorrhage with Amyloidosis (HCHWA), Icelandic Type in mutant form L68Q*

Abbrev: SAD, Sporadic Alzheimer’s disease; FAD, Familial Alzheimer’s disease; HD, Huntington’s disease; SBMA, Spinal and Bulbar Muscular Atrophy; SCA, Spinocerebellar Ataxia; DRPLA, Dentatorubral-Pallidolaysian Atrophy * See text for examples of CAA, cerebral angiopathy for the APP Dutch (E693Q), Flemish (A692G, Italian (E693K), Arctic (E693Q), or Iowa (D694N) mutations > (Figure 16-1).

Promoting Abx clearance to prevent intracellular build up: degradation via action of different proteases. Recent examples include insulin-degrading enzyme (IDE), neprilysin (endopeptidase 24.11), and cathepsins, or the stimulation of proteasome by resveratrol found in red wine (Marks et al., 1994; Carson and Turner, 2002; Hersh, 2003; Tanzi et al., 2004; Marambaud et al., 2005; Saito et al., 2005b).

1.2 APP Heterogeneity Alternative splicing of an 18-exon gene and posttranslational modifications account for the APP heterogeneity that likely alters their localization and processing. For example, APP695, the major neuronal isoform, lacks the Kunitz protease inhibitor (KPI) sequence or other domains coded by exons 7 and 8 present in APP751/770, but all these possess a C-terminal consensus sequence for endocytosis (> Figure 16-1). Other members of the superfamily include APP-like proteins (APLP1, 2), which lack the Ab sequence and show

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. Figure 16-2 Strategies for therapeutic intervention. > Table 16-1 for abbreviations

no link to AD (Coulson et al., 2000). APP knockouts (KO) remain viable but those of APLP-1 and -2 die shortly after birth, suggesting roles in development (Herms et al., 2004). Abx forms constitutively but its function, like that of APP, remains to be established. It has been suggested that its formation represents an unsuccessful response to stress. In the case of the precursor its wide distribution indicates its roles in general cell housekeeping, as seen from the presence of a heparinbinding domain for cell adhesion, in ion flux, or in kinase-mediated GMP signaling, etc. While its function is still a subject of debate, most studies focus on aberrant processing as a factor in amyloidogenesis. AD is multifactorial with probable cross talk between multiple genes and risk factors of which APP is one. Sporadic AD (SAD), which comprises approximately 95% of all dementia of the Alzheimer-type cases, occurs at 65 years and has no established genetic linkage other than ApoE alleles acting as a risk factor. This contrasts with FAD for approximately 5% of cases with dementia that occurs below 65 years, with linkage to APP or PS mutations. A general feature of FAD is an increase in the formation of Ab 1–42 (A42), which is more prone to fibrillogenesis than shorter forms. FAD facilitates the creation the creation of Tgs that overexpress mutant forms of APP/PS or the preparation of transfected cell models. FAD is also associated with mutations in PS, which are discussed more fully in subsequent sections (> Figure 16-1,


16

> Figure 16-9). Importance is attached to the ratio of A42/40, although there are exceptions. A recent Tg model that specifically increases only A42 without altering levels of precursor also results in amyloidogenesis (McGowan et al., 2005). The spectrum of APP metabolites varies with each Tg model. An example is shown for PDAPP (Val717-Phe), which surprisingly lacks C83 and C99 as major metabolites (Esh et al., 2005). These discoveries have stimulated considerable interest in the manner mutations alter APP processing and notably shift sites of cleavage to favor longer and more fibrillogenic products. This has led to some notable findings on protein turnover particularly for membrane-bound components. It has been noted frequently that many FAD mutations occur at or near sites of secretase processing that may alter secondary structure and processing (> Figure 16-1). Mutations at A21-D23 (Abx numbering, > Figure 16-1) for the most part are not associated with dementia, but induce cerebral amyloid angiopathy (CAA) (Tian et al., 2004). Thus, such mutations provide scope of developing other Tg models to investigate this phenomenon, especially since it is an unwanted side effect of anti-Abx immunotherapy (Sennvik et al., 2002; Herzig et al., 2004; Roher et al., 2004). Hopefully such models can be used to evaluate nonfibrilforming immunogens, Ab12–28Val18Pro, or nonspecific IgGs to attenuate amyloidogenesis without inducing CAA (Sadowski et al., 2004). In addition, CAA is also associated with transerythretin, gelsolin, mutated cystatin C, and ABr/ADn among a growing list of proteins of diverse structure within a broad group of amyloidoses (> Table 16-2). This emphasizes the deleterious effects of 3D structure as an etiological factor. Even in the case of APP, a recent study on isomerization mediated by prolyl isomerse showed that Pin-1 binding to pT688-P elevates A42 and is associated with the formation of SP and tangles by pathways replicated in part in a KO model (Pastorino et al., 2006).

1.3 Regulated Intramembrane Proteolysis Emerging evidence within the past 5 years established the existence of multimeric complexes mediating transmembrane (TM) cleavage not only for APP but also for other type-1 proteins. This recent discovery supersedes earlier findings that remain contradictory or inferential. This finding was made possible by the association of PS to FAD and its binding to nicastrin (NCT), a mammalian ortholog of Caenorhabditis elegans aph-2. This provided an impetus to screen nematode DNAs for other components critical for the cleavage of type-1 protein Notch, thereby releasing a notch intracellular domain (NICD) for signaling. This affirms the importance of earlier studies on FAD as a model to investigate regulated intramembrane proteolysis (RIP), now also relevant to similar conversions seen in SAD. > Table 16-1 indicates the current nomenclature and key references for ‘‘obligatory’’ components of the multimeric g-secretase complex (g-SC). g-SC has not yet been isolated in a homogenous form to establish the configuration of components or the manner of docking to substrate. Secretases are reviewed here in the order of their historical discovery with emphasis on enzymology as a basis to prepare useful clinical agents. In view of the large and growing literature, an attempt is made to summarize properties whenever possible in the form of lists, tables, and figures.

2

a-Secretases (K16L17 Cleavage) and the ADAM Gene Family

The brain lacks a specific KL-cleaving enzyme, but metalloproteases of the ADAM family share this property. These are a subgroup of metallozinc enzymes of the adamalysin/reprolysin superfamily with an HEXGHXXHGXXHD metallocenter (> Figure 16-3). Products include C-83 as a substrate for RIP, sAPPa with putative trophic properties, and a residual peptide P3 with no known function (> Figure 16-1) (Skovronsky et al., 2000a; Blacker et al., 2002).,

2.1 Characterization Interest in the purification of ADAMs stems from their potential roles in normal APP turnover to prevent amyloidogenesis. Human and murine ADAM-10 genes have 160 kb with 16 exons and lack specific

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. Figure 16-3 ADAM (TACE, ADAM-17) domains. Sites for alteration of activity (see text) ( , ) DIS, disintegrin; EGF, epidermal growth factor; Cyto, cytoplasmic tail. > Figure 16-1 for other symbols

polymorphisms that link to AD. Presence of RA response elements, Sp1 and USF, within a core promoter region located 508 to –300 from the ORF provide sites for regulating transcription (Prinzen et al., 2005). There are more than 30 cDNAs in human or other mammalian tissues but 17 actually code for enzymatic forms that play diverse roles in cellular function including growth, differentiation, cell fusion, and fertilization. Surprisingly, very little has been established for comparable roles in the CNS other than the nonspecific shedding of ectodomains from various proteins. These include conversion of proTNF-a to release the inflammatory 26-kDa (TNF-a) cytokine or the processing of ACE, Notch, Delta, Jagged, L-selectin, EGFR, IL-6R, APLP-1/2, prions, oxytocinase, interleukin receptors, and HER-4 trk (Schlondorff and Blobel, 1999; Brou et al., 2000; Mumm et al., 2000; Primakoff and Myles, 2000; Rio et al., 2000; Parkin et al., 2002; Becherer and Blobel, 2003; LaVoie and Selkoe, 2003; Seals and Courtneidge, 2003; Eggert et al., 2004). The relevance of many of these to dementia has not been fully established. Studies on structure–activity relationships for APP are scarce. A study on APP constructs in COS cells showed that KL cleavage was strongly influenced by upstream motifs (Sisodia, 1992) with similar conclusions for Fas/p75TNFR chimeras on expression in K293 cells, implying that secondary structure was a major determinant (Zampieri et al., 2005). The major motivating factor in purification has been studies directed toward synthesis of antiinflammatory agents. For this purpose, the processing sites of TNF provide small peptides including AcSPLAQAVRRSSR for rapid assay. A comparable APP sequence Ac-YHHQKLVFFA-NH2 provides a basis for seeking enzyme(s) with higher affinity for this substrate aided by chromogenic additions to distinguish different members of the superfamily: Mca-PLAQAV-Dpm-RSSSR or Mca-PLGLDpaAR for ADAM-17 and Dnp-PLGLWA-D-R for ADAM-9, etc. The use of different strategies, using transfected or other cell models, established ADAMs as putative a-secretases for KL cleavage. These include CHOAPP751 cells, an absence of conversion by null fibroblasts, or use of dominant negative ADAM mutants that lack the Zn2þ metallocenter (Buxbaum et al., 1998; Lammich et al., 1999).


16

2.2 Domain Structure and Therapeutic Implications TACE or ADAM-17 is a prototypic ADAM used to identify domains for drug interactions (> Figure 16-3) (Black et al., 1997; Moss et al., 1997; Buxbaum et al., 1998). Preferably, activation is more desirable to promote normal APP turnover, but other than the use of 4-aminophenylmercuric acetate (APMA) (Allinson et al., 2004), few studies direct attention toward the direct interaction with the catalytic center. While its mechanism requires further study, APMA promotes the conversion of epidermal growth factor HB-EGF and TNF-related protein TRANCE (Schlondorff et al., 2001). A potentially beneficial event may result from the conversion of inactive proenzyme via the conserved ‘‘furin’’ RX/R/K/R site and its coordination with the ‘‘cysteine switch’’ (> Figure 16-3). This has been difficult to confirm except indirectly by the action of the furin inhibitor a-1-PDX on transfected K293 cells that overexpress PC7 convertase (LopezPerez et al., 1999). As noted, most studies focus on antiinflammatory actions to prevent cytokine release by mechanisms mediated by the Zn2þ-catalytic center (Maskos et al., 1998). Even though AD has an inflammatory component, few studies exist on suppression as a therapeutic goal. Chelating reagents lack specificity but hydroxamates have some selectivity, with batimastat having >100-fold more potency for ADAM-17 or collagenase and marimastat having 4-fold more potency for APP (Parvathy et al., 1998; Newton and Decicco, 1999; Parkin et al., 2002)(> Figure 16-1). The modeling of ADAM S1/S3 pockets may provide better guidance for designing specific antiinflammatory agents (Manzetti et al., 2003), but relevance for APP processing again is questionable. Other domains as potential drug targets have not received interest in terms of therapy. These include EGF repeats, the cysteine-rich region, disintegrin (an integrin-binding domain), and variably sized cytoplasmic tails (Primakoff and Myles, 2000; Smith et al., 2002; Seals and Courtneidge, 2003). It has been proposed that the binding of disintegrin to substrate alters the conformation of the Zn2þ-catalytic center, as shown for APP in > Figure 16-4. Disintegrin has some homology to a 90-mer snake venom neprolysin (SVR) despite the absence of an RGD motif, which induces fatal bleeding on binding to GpIIb/IIIa integrin receptors (Seals and Courtneidge, 2003). Whether there is any relationship to angiopathy remains

. Figure 16-4 Changes in conformation on binding of DIS/EGF domains of ADAM to APP to promote a-secretase cleavage

477

478

16


undetermined. The cysteine-rich domain binds ‘‘syndecan,’’ a proteoglycan similar to one in SP, which coincidently also contains irADAM. This raises the possibility that proteoglycans or their metabolites are factors in signaling, similar to their roles in the CASK pathway (Skovronsky et al., 2001; Gatta et al., 2002; Bernstein et al., 2003; Schulz et al., 2003). Finally, the cytoplasmic tails contain sites for phosphorylation and may influence organelle trafficking on binding to SH2/SH3 SH3PX1 or GrB2 (endophilins) (Howard et al., 1999; DiazRodriguez et al., 2002). Evidently if ADAMs play important roles in amyloidogenesis as is increasingly clear from a number of pharmacological studies, then further studies on domain structure may be productive. The CNS, like other tissues, contains endogenous tissue inhibitors of metalloproteases, TIMP 1–4 with Mr 184–195 kDa, which have been associated with brain arteriovenous malformations (BAVM) or are involved in altering the viability of DOX-treated embryonic cortical cells (Hashimoto et al., 2003; Wetzel et al., 2003), but no studies exist in demonstrating their role in AD/FAD. However, it may be possible to design agents targeting certain ADAM family members based on TIMPs: ADAM-17 is selectively targeted by TIMP-3 whereas ADAM-10 is inhibited by both TIMP-1 and -3 (Lee et al., 2005).

2.3 Constitutive and Regulated APP Processing While phorbol esters promote the nonamyloidogenic pathway, ADAMs act indirectly without changing the phosphorylation of APP itself (Hung and Selkoe, 1994; Racchi et al., 1999). This suggests that they act on ancillary pathways mediated by specific synaptic kinases, for example PKCe, (Zhu et al., 2001) or on others including those mediated by PKA, MAP, TrkA, C-Jun, ERK, P13, GSK-3a, Cdk5, but data remain conflicting (Vincent, 2004). It is unknown which of several ADAMs act as a key secretase since candidates include: ADAM-9 (MDC, Meltrin-g and MCDP)(Roghani et al., 1999) ADAM-10 (Kuzbanian or KUZ, MADAM, Sup-17) and ADAM-17 (TACE) (see domain structure in > Figure 16-3). Which ADAM is paramount for APP processing is an unresolved issue owing to relative lack of specificity. This may ultimately be a question of localization and transport. Some studies point to ADAM-10 over -17 as in furin-deficient LoVo cells expressing proconvertase-7 (Lopez-Perez et al., 2001), in neural-like SH-SY5Y, or in kidney HEK 293 cells (Endres et al., 2003). However, RNAi in a human glioblastoma A172 cell model (Asai et al., 2003) or in embryonic ADAM-9 KO cells imply redundancy (Weskamp et al., 2002). ADAM deletion can induce fatal cardiac and CNS malformations, which limits use of null cells (Hartmann et al., 2002). Studies on heterozygous ADAM-10þ/ cells favor this member of the superfamily for APP conversion (Postina et al., 2004; Villanueva de la Torre et al., 2004). There remains the possibility for a fine balance between secretases competing for a limited pool of APP such that promoting ADAM activity is at the expense of aberrant processing. This is implied by the presence of ADAM in SP (Bernstein et al., 2003; Olsson et al., 2003) and the colocalization of ADAM mRNAs at sites for APP processing, but this is an unresolved issue in AD (Marcinkiewicz and Seidah, 2000).

2.4 Alternative a-Secretases It is not known if tissues contain an ADAM with a high affinity for APP, which can be used as a potential drug target. Earlier studies that showed yeast Kex-mRNA acting as a bait to identify a mammalian prohormone-converting enzyme homolog can provide a precedent. Yeast contains unique Asp proteases (Yap3 and McK7 or yapsins), which restore defective vacuolar transport in mutants and also act on APPsubstrate Ac-RE(Edans)VHHQKLVPFK-(dabcyl) at the appropriate a-secretase site (Zhang et al., 1997; Komano et al., 1998; Greenfield et al., 1999).


16

2.5 Therapeutics PKC activation: (phorbol esters, GF109203X) (Yeon et al., 2001; Zhu et al., 2001). Atorvastin, a statin, acts indirectly to promote a-secretases without changes in PKC and ERK in Tgs or transfected cell models (Parvathy et al., 2004). The pathways remain to be mapped but activation may be mediated via CD9-tetraspanins present in lipid rafts (Moss and Bartsch, 2004). Pituitary adenylate cyclase-activity polypeptide (PACAP) promoted a-APPs via activation of a PAC-1 receptor with inhibition by PACAP (6-38) acting as an antagonist. This results from PK inhibition mediated by MAPkinases erk-1/2 and phosphatidylinositol-3 (Kojro et al., 2006). Cholinergic activation by use of muscarinic agents: AF150, -102B, and N277B (Fisher et al., 2002; Fisher et al., 2003) Cholesterol-lowering drugs: (methyl b-cyclodextrin, statins, BM15.766, TV3326, filipin). In view of their general use for treatment of hypertension, there has been considerable interest and controversy whether cholesterol-lowering drugs are also beneficial in reducing the risk of AD. These may act on secretase binding to GPI anchors in lipid rafts to potentially alter APP turnover (Kojro et al., 2001; Runz et al., 2002; Parkin et al., 2004). This has prompted the use of statins to lower blood cholesterol for the treatment of dementia (Parvathy et al., 2004). Inhibition of HMG-CoA synthase prevents the formation of mevalonate within the cholesterol biosynthetic pathway and isoprenoid units acting as kinase modulators. Concomitant changes in membrane fluidity may alter interaction between secretases and substrates within the innermembrane leaflets or lipid rafts. Statin proforms Simovastin or Lovastatin penetrate the CNS barrier for esterase activation in Tgs (Mark Burns, personal communication, NKI). BM15.766 acts downstream to alter cholesterol reductase (Refolo et al., 2001). A study on artificial unilamellar vesicles containing BACE demonstrates activation by cerebrosides, glycerophospholipids, and sterols (Kalvodova et al., 2005). Pharmacological agents: Amphiphiles, imipramine, MAO-B inhibitors (Rasagaline, N-propargyl-(IR)aminoindan, Ladostigil) (Yogev-Falach et al., 2003; Youdim et al., 2005). Donepezil, an AchE1 inhibitor, impacts trafficking of ADAM-10 in SH-5Y5Y cells (Zimmermann et al., 2004). There is considerable scope for this idea, even for the use of diet components such as epigallocatechin-3-gallate found in green tea (Rezai-Zadeh et al., 2005). NSAIDs: (Nonsteroidal antiinflammatory drugs). Some NSAIDs suppress Abx formation (flurbiprofen, sulindac sulfide), but others are inactive (Avramovich et al., 2002). These may act preferentially on the allosteric site(s) of the g-SC (> Sect. 4.5, > Table 16-4) (Gasparini et al., 2004) or activate peroxisome proliferator-activated receptor g(PDAR-g) with an impact on BACE mRNA expression (Sastre et al., 2006). Immunotherapy: A novel but plausible approach is the use of ADAM IgGs or fragments (hk14, C23.5) with putative enzymatic properties (Rangan et al., 2003; Liu et al., 2004). Miscellaneous: APLP-1 reduces APP shedding via a common LRP-mediated pathway and a shared NPTY C-terminal motif (Neumann et al., 2006).

3

b-Secretases (BACE-1 and -2)

Secretases convert APP at the Met1-Asp1 (b site) for the secretion of sAPPb and the formation of a cellassociated C-99 as shown in > Figure 16-1, hence the term b-APP-converting enzymes (BACE, b-secretases) (called also memapsins or Asp proteases) (> Table 16-1). Cleavage upstream at Tyr10-Glu11 (b1 site) yields N-truncated CTFs as a second metabolite, detectable in tissue extracts (> Figure 16-1). Two forms exist: BACE-1 and -2 coded by different genes with BACE-2 associated with DS and acting also at F19F20 (Saunders et al., 1999; Acquati et al., 2000; Yan et al., 2001b; Fluhrer et al., 2002; Motonaga et al., 2002; Barbiero et al., 2003). BACE-1 and -2 share an 50% homology, but the enzymes have divergent N- and C-termini. These may act as factors contributing to different histopathologies seen in AD/DS. The two distinct forms may account for the different progression of AD and DS and for the deposition of SP/CD (Willem et al., 2004).

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480

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3.1 Domain Organization and Implications for Therapy The deduced sequence of BACE-1 provides information on the domain structure (> Figure 16-5). It indicates prepro- and propeptides, a single TM contiguous to a loop region, and a conserved D*S/TGS/T catalytic center (Yan et al., 2001a; Schmechel et al., 2004; Shiba et al., 2004; Westmeyer et al., 2004). Six . Figure 16-5 BACE-1 domains. Structure and potency of two typical site-specific inhibitors indicated. The disulfide sequence at 330–380 (bold) is required for activity of the consensus catalytic center. For other abbreviations and symbols > Figures 16-1 and 16-3

cysteines form three disulfide bridges, but only the pair at C330/C380 is essential for catalysis (Fischer et al., 2002). Alternative splicing accounts for 476, 457, and 432 isoforms but none of these link specifically to neurodegeneration (Tanahashi and Tabira, 2001). BACE is rate limiting for the formation of C-99 and thus presents an attractive target for therapy to reduce amyloidogenesis (> Figure 16-2). Targets include furin conversion of proenzyme (Bennett et al., 2000) or conversion at motifs to reduce trafficking to sites for APP processing. Those shown in > Figure 16-5 include an acidic dileucine (ACDL) motif that binds to the VHS (Vps-27, Hrs, and STAM) domain of GGA (Golgi-localized g-ear-containing AR-binding) or to a Ser498 for phosphorylation (He et al., 2005), or to PAR-4 (prostate apoptosis response-4), a leucine zipper protein on the C terminus (Xie and Guo, 2005). Other promising targets include the transcriptional elements P1/2, CREB, GC box, NF-kB, and STAT 1 present on the 30.6-kb promoter region of hBACE-1 (chromosome 11 band 23.2-11q23.3) (Christensen et al., 2004; Ge et al., 2004; Lammich et al., 2004; Sambamurti et al., 2004). The expression of the BACE-1 gene is low in vivo and its control is not well-defined. A recent analysis identifies six upstream AUG sites in the 50 -UTR of human mRNA: the first pair is essential in the promoter region, while the fourth acts as a translation initiation codon, with deletion of mutation enhancing expression and requiring an intact eIF4G (Zhou and Song, 2006).

3.1.1 Characterization BACE was the first specific secretase to be purified and shown to be an unusual Asp protease different from classical forms. The methods used followed standard procedures of protein separation or DNA screening (Hussain et al., 1999; Sinha et al., 1999; Vassar et al., 1999; Yan et al., 1999; Lin et al., 2000). An example of the first includes affinity chromatography on columns coupled to the inhibitor KTEEISEVNLstatinVAEF to


16

purify human brain BACE (Sinha et al., 1999). The second includes screening of DNA libraries to identify similar enzymes having alternative nomenclatures: asp-2/memapsin-2 and asp-1/memapsin-1 identical to BACE-1 and BACE-2, respectively. Screens also identified other Asp proteases not active toward APP (Asp3/4, napsins 1 and 2) (Tatnell et al., 1998; Yan et al., 1999; Schauer-Vukasinovic et al., 2000). All studies show higher rates for processing at the b-site of APP or surrogate peptides bearing the Swedish NL mutation (> Figure 16-1).

3.2 Specificity While studies on specificity are incomplete, it is apparent that the term BACE is a misnomer since it acts also on other proteins relevant to immunology and cell signaling (> Table 16-3). This will have to be taken into consideration in developing inhibitors to target solely APP (Lin et al., 2000; Gruninger-Leitch et al., 2002; Turner III et al., 2002; Turner III et al., 2005). Other proteins include P-selectin glycoprotein ligand-1 (PSGL-1) (Lichtenthaler et al., 2003), 2,6 sialtransferase (ST6Gal 1) (Kitazume et al., 2001; Kitazume et al., 2003), APLP1/2 (Eggert et al., 2004; Li and Sudhof, 2004; Pastorino et al., 2004), the voltage-gated sodium channel b subunits 1–4 (Wong et al., 2005), and the low-density lipoprotein receptor (von Arnim et al., 2005). Interestingly Abx itself serves as a substrate indicating dual roles for BACE in both its formation and clearance (Lin et al., 2000; Shi et al., 2003). Clues on the design of site-specific inhibitors are provided by the use of peptides that show a preference for cleavage at P4-P40 with some long-range influence by residues at P5-P7 or from crystallography of an OM03-4/enzyme–inhibitor complex, indicating the existence of an a-helical domain and a cleft or flap region for accommodating agents (Hong et al., 2000; Hong and Tang, 2004; Xiong et al., 2004; Turner III et al., 2005). Mutagenesis to insert disruptive Pro or other residues points to conformation as a key factor in the cleavage of constructs and offers the possibility to target preferentially the b- and b0 sites (Qahwash et al., 2004).

3.3 Pharmacology BACE as a rate-limiting event for Abx offers a potential target to reduce amyloid burden especially since it is nonessential for development, as seen from KO studies (Roberds et al., 2001). There are reports on behavioral changes in multiple Tg models with BACE-1 KO ablating temporal memory deficits mediated by the hippocampus, conforming to the view that this pathway may be amenable to therapeutic intervention (Ohno et al., 2006). Multiple studies show increase not only in AD but also in ischemia and CAA, as the result of ageing (Rossner et al., 2001; Fukumoto et al., 2002; Holsinger et al., 2002; Hartlage-Rubsamen et al., 2003; Hong et al., 2003; Li et al., 2003a; Fukumoto et al., 2004; Li et al., 2004; Sennvik et al., 2004; Wen et al., 2004; Rossner et al., 2005). Such increases seen in Tg models (Chiocco et al., 2004; Willem et al., 2004) are without a concomitant change in transcription, implicating a role for ancillary pathways (Roberds et al., 2001; Harrison et al., 2003; Sant’Angelo et al., 2003). There also is a report on high expression using a Thy1 promoter in Tgs to induce a form of neurodegeneration without reduction in Abx (Rockenstein et al., 2005). Despite these paradoxes and the lack of specificity, there have been a large number of attempts to modulate activity as follows: Trafficking: Methyl-b-cyclodextrin or leptin has been used to target the trafficking of BACE within membranes or its attachment to lipid rafts via GPI-anchored proteins (Cordy et al., 2003; Marlow et al., 2003; Scholefield et al., 2003; Fewlass et al., 2004; Huang et al., 2004). Brefeldin or monesin to alter targeting to Golgi complex/ER or propeptide processing in cell models, or agents to promote the ubiquitin-proteasome pathway for enzyme inactivation during transport, have also been used (Bennett et al., 2000; Qing et al., 2004). Other targets to influence transport as a rate-limiting factor include: 1. ACDL for binding to the VHS domain of the adaptor protein GGA (Shiba et al., 2004) 2. Phosphorylation of Ser498 in the C terminus of the cytoplasmic domain to influence internalization (Walter et al., 2001; von Arnim et al., 2004) 3. Transport mediation by Nogo (He et al., 2004) or type II BRI3 (Wickham et al., 2005)

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. Table 16-3 Specificity of BACE 1/2 1/2

Substrate, cleavage Ab40/42 –GSNKGAIIGL34↑MVGGVV40IA42

Comments g-SC dependent

1/2

APP at b site –KTEEISEVKM↑D1AEFRHDS–

1/2

APP at b0 site –DAEFRHDSGY10↑11EVHHQ–

Increases with the FAD double Swedish mutation NL↑D Rodents and humans

1

Abz-VKM↑DAE-ethylene diamine-Dnp Abz-VNL↑DAE-ethylene diamine-Dnp SEVKM↑DAEFR SEVNL↑DAEFR VVEVDA↑AVTP 7-methoxycoumarin-4-acetyl-EVNL↑DAEF-KDnp-COOH Ac-EIDL↑MVLDWHD-K-Dnp

1

1 1 1

a2,6-Sialtransferase –31DYEALTL37↑QAKEFQMPKSQE49–

1

Voltage-gated sodium channel b-subunits 1–4 b1 –NTSVVKKIHL144↑145EVVDKANRDM– b2 –RHRGHGKIYL144↑145GVLLEVPPER– b31 –NVSREFEF128↑129EAHRPFV135– b32 –136KTTRLIPLRVTEEAGEDF153↑154TSVVSE– b4 –DLNNSATIFL149↑150QVVDKLEKVD–

1

P-Selectin glycoprotein ligand-1 (CD162)–KGIPMAASNL303↑S304VNYPVGA–

1

APLP2 VKEMVIDETL659*D660VKEMIFNAE–APLP –KVNAS553*554VPRGFPFHSSEIQR567*568DELA–

2

Ab1–28 –DAEFRHDSGY10↑11EVHHQKLVF19↑F20 AEDVGSNK– BACE-2 prodomain –HADGLAL62↑63 ALEPALA– APP within b-peptide –DAEFRHDSGYEVHHQKLVF19↑F20 AEDVGSNK–

2 2

Synthetic surrogates Prefers L at P1, polar/acidic 0 residues at P2, P2 , hydrophobic at P3 (best: V)

P1 site accommodates bulky, hydrophobic (L, F, M, Y); also at P30; type-2 protein Cleavage may affect neurochemical and behavioral functions relevant to altered elecrtrochemical balance. Similar to APP, sequential processing, first by b-secretase then g-secretase Homology to APPSW sequence; dependent on presence of transmembrane domain 17 residues downstream; Type-1 protein Postulated APLP 1/2 cleavage point based on homology to APP; APLP1 cleavage within domain bounded by open arrows

Autocatalysis/activation Occurs in Golgi complex and later secretory compartments, prevalent in Down’s syndrome

Reference Fluhrer et al. (2003) Sinha et al. (1999); Farzan et al. (2000) Qahwash et al. (2004) Andrau et al. (2003) GruningerLeitch et al. (2002) Marcinkeviciene et al. (2001) Toulokhonova et al. (2003) Kitazume et al. (2003) Wong et al. (2005)

Lichtenthaler et al. (2003)

Li and Sudhof (2004)

Farzan et al. (2000) Hussain et al. (2001) Fluhrer et al. (2002)


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4.

4

Neuronal sorting protein-related receptor sorLA as a trafficking receptor to prevent BACE–APP interaction (Spoelgen et al., 2006) 5. Using dileucine motif (L499-L500) to direct the enzyme to a late endosomal/lysosomal compartment for degradation: lysosomal inhibitors chloroquine and ammonium chloride lead to accumulation of enzyme (Koh et al., 2005) Transcriptional regulation: By using SP1 and AP2 or others within the 30 - or 50 - untranslated domains as mentioned above (Sambamurti et al., 2004) Using inhibitory substrate analogs with statins and isotere insertions (Roggo, 2002): Delivery to cells may be facilitated by viral vectors or surrogate carrier peptides such as HIV Tat46–57 sequence (GYGRKKRRQRRR) (Wender et al., 2000; Cronshaw et al., 2003; Chang et al., 2004). Transitionstate analog KMI-429 inhibits activity in cultured cells or on direct injection into the hypothalamus of Tg models (Asai et al., 2006) Immunology: Utilizing sequences adjacent to b sites to raise antisera (Arbel et al., 2005; Paganetti et al., 2005) Inhibitory nucleic acids: Antisense oligos, RNAi, etc. (Kao et al., 2004; Nawrot, 2004). Lentiviral delivery of BACE-1 siRNA reduces APP cleavage in TgAPP mice (Singer et al., 2005) supporting the view that increased BACE-1 expression elicits neurodegeneration in such models (Rockenstein et al., 2005) Miscellaneous: Ad hoc screening of plant or fungal extracts has yielded talsaclidine, hispidin, tenuigenin (Jia et al., 2004; Zuchner et al., 2004), and other potential lead compounds including benzofuran and analogs (Espeseth et al., 2005)

g-Secretase Complex

4.1 Clarifying g-SC as a Multimeric Complex Studies on g-SC are the least satisfying due to the nonavailability of purified forms. A major finding that supersedes earlier attempts to resolve mechanisms of Abx formation was the identification of NCT as an essential binding partner for PS. This introduced the concept of multimeric complexes and the discovery of other binding partners. NCT is a mammalian ortholog of nematode aph-2, which previously was found to facilitate Notch processing and signaling. Subsequent screening of nematode DNAs led to the identification of other components of the multimeric complex shown in > Table 16-4 that minimally are required for Abx production although this does not exclude others that play ancillary roles. The chronology for this major breakthrough is as follows: NCT (a C. elegans aph-2 homolog) is a binding partner for PS (Yu et al., 2000) Nematode DNA screening to identify aph-1 and pen-2 as additional binding partners (Francis et al., 2002; Goutte et al., 2002) Confirmation for multimeric complexes in human brain extracts by immunoprecipitation (Kimberly et al., 2003b) Reconstitution in yeast or other cell models with suppression by RNAi or use of putative inhibitors of PS-mediated events (Francis et al., 2002; Goutte et al., 2002; Edbauer et al., 2003; Farmery et al., 2003; Fraering et al., 2004; Prokop et al., 2004; Wrigley et al., 2005; Xie et al., 2005a; Zhang et al., 2005a)

4.2 Stepwise Assembly and Maturation of Complexes The use of RNAi and other methods established the step-wise assembly of mature active complexes (De Strooper, 2003; Iwatsubo, 2004; Steiner, 2004), shown in > Figure 16-6. This includes binding of aph-1 and NCT to form an inactive dimer for binding to PS. The overall complex has 18 TM but this may be subject to revision if other components are identified.

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. Table 16-4 Properties and structural determinants of obligate g-SC components Component PS1 TMP21b PS2

Gene locus 14q24.3 14q24.3 1q31-q42

NCT

1q22-q23

Aph-1a

1p36.13

Aph-1b

15q22.2

Pen-2

1q31-q42

Residues/Mr 467-mer 219-mer 448-mer FL: 43 kDa NTF: 27/CTF: 17 kDa 709-mer 80 kDa nonglycosylated 110 kDa immature 125–150 kDa mature aL: 265-mer aS: 247-mer b: 257-mer 30 kDa 101-mer

TM 7–9a 1

Determinant CTF: PAL, PALIY; NTF: Tyr288 C-terminal K215KLIE Asp257/385

1

D336YIGS

7

GxxxG (TM 4)

2

D90YLSF (C-terminal)

a

The number of TM domains is dependent on different algorithms; > Sect. 5.1, > Figs. 16-6 and 16-9 Not essential for formation of a tetrameric complex, but may direct PS-mediated g-SC cleavage of APP to preferentially release Abx, rather than other metabolites (Chen 2006); > Sect. 4.3 and 4.4.1 b

. Figure 16-6 Stepwise assembly of g-SC. (1) Binding of NCT ( )to aph-1 ( ) to form an inactive LMW complex; (2) Integration of PS holoprotein ( ) to form a stable HMW intermediate; (3,4) Addition of pen-2 ( ) to induce PS endoproteolysis to form an active g-SC; (5) It has been postulated the active complex on docking to substrates mediates RIP. Cylinders denote putative TM domains, (- -) and represent N-, C-terminal domains of NCT, which are differentially glycosylated ( ) (see text). > Figure 16-9 for position of key Asp residues ( ) that form a putative catalytic center ( )


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Addition of pen-2 to the complex is thought to trigger PS cleavage to form a catalytically active heterodimer for subsequent docking to and processing of C-99. This scenario is supported by pen-2 knockdown in transfected cells and by the critical role for a C-terminal motif (Prokop et al., 2004). There is no evidence that pen-2 triggers a cryptic protease within the complex to initiate proteolysis of labile PS. Maturation of the complex parallels an increase in NCT glycosylation from a 110 to 150 kDa form, which is speculated to provide a scaffold, as shown in > Figure 16-6, for the assembly of the complex (LaVoie et al., 2003; Luo et al., 2003; Nyabi et al., 2003; Takasugi et al., 2003; Kaether et al., 2004; Niimura et al., 2005) during transport from ER to Golgi complex, plasma membrane, or even mitochondria (Hansson et al., 2004; Capell et al., 2005; Chyung et al., 2005; Hansson et al., 2005; Nakaya et al., 2005). This may explain earlier anomalies in the distribution of PS relative to the processing of substrates termed ‘‘the spatial paradox.’’ Recent studies shed light on this topic since complexes occur within membrane microdomains or lipid rafts for delivery via endosomal or autophagic pathways mediated by rab 6/8 or GDI (GTP dissociation inhibitor) (Torp et al., 2003; Wada et al., 2003; Bergamini et al., 2004; Pasternak et al., 2004; Vetrivel et al., 2004; Yu et al., 2004). g-SC in nonraft membranes facilitates TM cleavage during embryogenesis, but translocation on maturation to raft microdomains differentially promotes C-99, rather than Notch or Jagged. Therefore, agents targeting enzyme localized in rafts provide a method to specifically target amyloidogenesis (Vetrivel et al., 2005). Alternatively, recent studies by Dewji et al. (Dewji et al., 2006) raise new questions on APP processing within a single cell or following surface interaction with other cells containing g-SC obligatory components via a lysosomal pathway. They demonstrate that on coculturing APP-null E18 hippocampal neurons with PS1/2-null ES cells transfected with hAPP results in internalization via a lysosomal pathway sensitive to impermeable N-termini of PS-1 or -2, thus generating Ab. This is consistent with the placement of the N terminus of PS extracellularly in the proposed 7-TM but not the 8-TM domain model for PS to account for surface interference by these peptides (> Sect. 5.1, ˚ interacting with lipid > Figure 16-9). A 3D-EM study suggests the complex exists as a cylinder 20–40 A bilayers (Lazarov et al., 2006), and the forms present on the plasma membrane are functionally distinct from those associated with endosomes (Fukumori et al., 2006). Assembly of variable isoforms for obligatory components with labile or stable PS may account for divergent specificities of complexes (Berezovska et al., 2003; Ramdya et al., 2003). For example, integration of biostable PSDEx9 mutant promotes A42 release (Prokop et al., 2004), whereas random mutagenesis results in biostable PS forms R278I or L435H with A43 release (Nakaya et al., 2005). It follows from these examples that PS proteolysis is not critical for TM cleavage. This is confirmed on transfection of these mutants in PS/ MEF cells, which remain sensitive to typical inhibitors MW167 and the NSAID sulindac sulfide. Studies using PS-deficient blastocytes (BD) indicate PS-independent g-SC sensitive to pepstatin A preferentially forming A42, which could indicate an alternative aspartyl protease activity at pH 6.0 (Lai et al., 2006).

4.3 Isolation of g-SC Detergents CHAPS, CHAPSO, or Brij 35 provide cell-free extracts for assays using C-99 or other TM proteins or their surrogates Nma-GGVVIATV-K, Nma-GCGVLL-K(Dnp), C-99-GV, NotchdE, and NecnGV (Dyrks et al., 1993; Capell et al., 1998; Yu et al., 1998; Li et al., 2000a; Francis et al., 2002; Beher et al., 2003; Farmery et al., 2003; Kimberly et al., 2003a; Carter et al., 2004; Gu et al., 2004; Gupta-Rossi et al., 2004; Wrigley et al., 2005). Detergent extraction may disrupt complexes on plasma membrane surfaces and recognition of bound or soluble recombinant substrates such as APP (Marks et al., 1995; Chyung et al., 2005). Localization remains to be defined, but could involve association with late autophagic vacuoles (AV) that contain APP, C-99, and several obligatory components of the g-SC (NCT, PS1) (Yu et al., 2005). Isolation of complexes from normal human brain points to nonmutated PS also as a key player in RIP (Farmery et al., 2003). Moreover, reconstitution in Saccharomyces cerevisiae lacking g-SC or in animal and insect cells affirm a critical role for all four components, indicated by the hydrolysis of constructs C1–55GAL4 and C-100-His6APP (Fraering et al., 2004; Niimura et al., 2005). Recent procedures involve the isolation of Flag-pen-2, aph1a2-HA, and NCT-GST from transfected CHO expressing hPS1 for affinity

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purification with III-C and capture by anti-Flag2 and GSH chromatography. This yields a 160-fold enrichment and a Mr of 400 kDa (Fraering et al., 2004), consistent for a tetramer or a simple multiple of the four components (Cervantes et al., 2004; Evin et al., 2005). Currently, no method claims homogeneity, and extracts contain other components believed to play ancillary roles in catalysis/trafficking by mechanisms that remain to be defined (> Table 16-5). This was confirmed for an 650-kDa complex purified from CHAPSO extracts of blastocytes expressing PS by isolation of TMP21, a member of the p24

. Table 16-5 Minor Components of g-SCa Component Ca2þ-binding proteins: Calsenins (DREAM-KchIP3)

CD147 ARD Cadherins and b-catenins bcl-2-related proteins Caveolin-3 Syntaxin PS1-374 isoform, CD92, prohibitin, transferrin receptor, IgG, ADP/ATP carrier protein–nucleotide-binding site, calmyrin, ubiquilin, PAMP/PARL

Comment and key references PS-binding protein: transcription factor regulating dynorphin function (Buxbaum, 2004), b-subunit of Kv channels (Lilliehook et al., 2003). Elevated in AD or neurons and astrocytes from Tgs; reduction in expression protects cells from Abx toxicity (Jo et al., 2004; Jo et al., 2005) Downregulates Ab in complexes from HeLa cell extracts (Zhou et al., 2005) Armadillo repeat proteins, P0071, B6P-plakophilin (Levesque et al., 1999) Signaling and transcriptional activation (Yu et al., 1998; Marambaud and Robakis, 2005) Cell death modulator (Alberici et al., 1999) Nishiyama et al. (1999) Smith et al. (2000) Miscellaneous components identified by i.p. of multimeric complexes (Mah et al., 2000; Zhu et al., 2004; Fraering et al., 2005)

It is not established if these are essential for g-SC, play ancillary roles, or represent contamination depending on the tissues and methods for extraction

a

family associated with sorting and concentration of secretory proteins (> Table 16-4 for size, genetic locus, and a major determinant) (Blum et al., 1996; Chen et al., 2006). Flag-tagged TMP21 immunoprecipitates other obligatory g-SC components from complexes of mouse brain, SH-SY5Y, and HEK cells. Importantly, siRNA suppression increases Abx release without effect on amyloid intracellular domain, notch intracellular domain, or CICD (cadherin intracellular domain), indicating a specific positive control of g- but not e-cleavage, thereby implicating a hitherto unsuspected role for PS in the differential cleavage of APP. This provides an avenue to develop agents to specifically alter amyloidogenesis. While TMP21 is not essential for the assembly of the tetrameric g-SC complex itself, the data imply a regulatory role in PS-directed amyloidogenesis. Loss of activity on treatment with SDS or with a change in pH is consistent with the multimeric status of complexes. Suppression by the agents 31-IIIC, DAPT, sulindac sulfide, L-685,458, pepstatin A, or difluoro alcohol 1 conforms to the known properties of g-SC (Evin et al., 2005). Relevant to the issue of other cell components are roles for brain lipids phosphatidylcholine, phosphatidyl ethanolamine, sphingomyelin, and cholesterol that alter activity (Fraering et al., 2004; Grimm et al., 2005; Wrigley et al., 2005). Statins as inhibitors of 3-hydroxy-3-methylglutaryl-CoA-reductase decrease C-99 processing in APP-transfected human neuroblastoma SH-SY5Y cells, by altering membrane cholesterol and fluidity or by binding of g-SC in lipid rafts (Urano et al., 2005).


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4.4 Specificity g-SC is nonspecific and acts on a variety of type-1 proteins to release ICDs of relevance to signaling or in the case of APP to pathology (> Figure 16-7). This may reflect a single protease with relaxed specificity . Figure 16-7 Release of intracellular domains (ICDs) and cell signaling following transport to the nucleus (----). Enzymes for sequential processing of holoproteins are indicated. In the case of cadherins, proteosomal degradation of CTF2 (C-terminal fragment-2 bound to CBP) and CREB-binding protein suppresses signaling. Multiple repeat domains in Notch (with copy number). EGF, epidermal growth factor (23); ANK, ankyrin (7); LN, Lin-12/Notch (3); MMP, matrix metalloproteases; FE65/Tip 60-nuclear transport proteins; CSL, CBF1/suppressor of hairless/Lag1; CREB, cAMP-dependent response element-binding protein; UPR, unfolding protein response; sensor of IRE-1 recognizes decrease of calcium stores in ER resulting in ectodomain shedding and g-SC cleavage, K/N, domain with both kinase and nuclease activities that splices XBP(X-box binding protein) mRNA to result in transcription of GRP78 (glucose-regulated protein), 78 kDa, a major ER chaperone; ♦, phosphorylation site. Other abbreviations and comments > Sect. 4.4. Adapted from Marambaud and Robakis (2005)

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acting sequentially, although existence of separate proteases cannot be excluded without purification (Zhao et al., 2005). There is a growing list of substrates and undoubtedly others will follow (> Figures 16-7 and > 16-8). . Figure 16-8 Cleavage of type-1 membrane proteins by g-SC. Nomenclature identifies proposed sites for intramembranal cleavage. In the case of APP, sequential processing of C terminally extended intermediates may precede Abx release. The epsilon site for APP (↑) resembles site-3 cleavage for Notch (> Figure 16-7). Boxes represent the TM domain

Notch: Sequential processing similar to other type-1 proteins with a C-terminal stub acting as a receptor (Notch-r). Unlike APP, it requires coupling to ligands to initiate downstream processing, releasing NICD for translocation to nuclei, which upregulates Drosophila suppressor of hairless (SuH) and the LAG-signaling pathway (Moehlmann et al., 2002; Roncarati et al., 2002). Four isoforms have been identified containing multiple EGF/ankyrin repeats and with KUZ/furin (sites 1, 2) for proenzyme conversion and one for NICD release mediated by g-SC (e or site-3). Notch-3 mutations are associated with cerebral angiopathy (CADASIL) and a mutation in its ligand Jagged is associated with Alagille syndrome, a mild form of mental retardation. This provides scope for investigating multiple ligands or inhibitors to signaling pathways that may be relevant to pathology in mammals (Delta1, Jagged1/2, numb), Drosophila (Delta, Serrate), and C. elegans (LAG-2, APX-1) (Sestan et al., 1999; Ikeuchi and Sisodia, 2003; LaVoie and Selkoe, 2003; Six et al., 2003). In addition to cleavage at the interface of the cytosol/ TM region to release NICD during Notch signaling (site-3 or e-like), studies have also identified a novel series of TM cleavages, releasing additional Nb products. These products are increased in presence of PS mutants and may be analogous to g-cleavages, which release Ab40 and Ab42 of APP (Okochi et al., 2006) APLP: Homologs APLP1 and 2 are members of the APP superfamily but lack the fibril-forming domain (> Sect. 1.2). APLP-2 is cleaved by g-SC to form an ICD capable of interacting with Fe65 (Walsh et al., 2003; Eggert et al., 2004)


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E-cadherin: A cell adhesion molecule that cleaves at Phe Leu751Arg, releasing an ICD (CTF-2) for translocation to nuclei, and altering catenin-Wnt pathway signaling (Marambaud et al., 2002) or repressing CBP/CREB transcription (Marambaud et al., 2003) Tyrosine kinase ErbB4: Processing to form an 80-kDa cytoplasmic transactivator EICD (Ni et al., 2001; Lee et al., 2002; Lai and Feng, 2004) Colony stimulating factor (CSF-1): Processing to release its CTF, suppressed by g-SC inhibitors, implying roles in regulation of PMA-mediated events (Wilhelmsen and van der Geer, 2004) LRP: Formation of fragments for binding to the Tip60/Fe65 complex for translocation of the LRP-ICD to nuclei where it may compete for Fe65/Tip60/APP AICD for transcriptional activation or affect APP processing (Fortini, 2002; Kinoshita et al., 2003; Yoon et al., 2005). LRP potentially competes with PS1 to displace APP within the complex (Lleo et al., 2005) CD44: Following the removal of ectodomain g-SC cleavage occurs at two sites, one resembling b-like in APP and one resembling Notch site-3, resulting in ICD that activates transcription mediated through the TPA-responsive element (Lammich et al., 2002; Murakami et al., 2003). A recent study indicates a role for CD44 in phagocytosis of macrophages containing g-SC obligatory components (Jutras et al., 2005) Ire1a: Associated with UPR (Unfolding Protein Response) (> Figure 16-7) or ATF6 (a type-2 protein associated with Ca2þ-ER stress) (Haze et al., 1999; Niwa et al., 1999), and decrease in the UPR chaperone GRP78, which is found coincidentally also to be decreased in AD brain (Katayama et al., 1999; Yu et al., 1999) Nectin-Ia: Processing by g-SC and roles in synaptogenesis (Kim et al., 2002) GluR3: Cleaves at L585-G586 to form GluR3b and interactions with GluR3a formed by cleavage in the cytoplasmic loop upstream at E570-P571 contribute to receptor diversity and specialization (Meyer et al., 2003) Megalin: A member of the LRP gene family localized to the kidney proximal tubule, upon PKC-mediated ectodomain shedding, its CTF acts as substrate for g-SC that may impact a signaling pathway (Zou et al., 2004) P75NTR: Cleavage of its CTF at a site more topologically related to amyloid formation than the e site-3 of Notch, forms a death receptor containing a death domain motif (NRADD) relevant to cell viability (Jung et al., 2003; Gowrishankar et al., 2004) IFNaR2 (subunit of type I IFN receptor): PMC-mediated sequential cleavage by g-SC to form a cellassociated product and ICD that appears to downregulate transcription of IFN-stimulated response elements (ISREs) (Saleh et al., 2004) Voltage-gated sodium channel b-2 subunit: A bifunctional member of the IgCAM superfamily that, following shedding of ectodomain by ADAM-10 or BACE, releases a 12-kDa ICD. Inhibition by DAPT or knockdown by PS RNAi results in diminished capacity for cell–cell adhesion or migration (Wong et al., 2005; Kim et al., 2005a) Growth hormone receptor (GRH): After ectodomain removal by TACE, processing of the CTF by g-SC produces an ICD for translocation to the nuclei, which potentially recruit other signaling pathways in adipocytes (Cowan et al., 2005)

4.4.1 AICD and Cell Signaling A product formed on downstream cleavage at L49V50 of C-99/C-83 of APP or by processing of APLP1/2 yields a sequence highly conserved in different species referred to as AICD (> Figures 16-1 and > 16-7). Formation is analogous to processing at site-3 (e) cleavage of Notch to release NICD, leading to interest in a role for this product in signaling (Kinoshita et al., 2002; Funamoto et al., 2004; > Figures 16-7 and > 16-8 for multiple sites and nomenclature for Notch cleavages by endoproteolysis). Earlier, AICD was not detected owing to extreme lability, due to rapid breakdown by IDE among others. Indeed, detection required the presence of a cocktail of protease inhibitors (Gu et al., 2001; Pinnix et al., 2001; Sastre et al., 2001; Yu et al., 2001a; Edbauer et al., 2002a; Chang et al., 2003).

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The absence of the bridge sequence between Ab and AICD (Ab41/43–49) may be due to lability, although sequential processing of intermediates cannot be excluded. Formation of AICD itself is another example of RIP by processing of APP. A recent study showed that blocking both g- and e-sites (> Figure 16-8), e.g., on insertion of Trp residues, apparently decreases Abx, indicating dependence of RIP on initial sites for processing (Sato et al., 2003; Sato et al., 2005). APP-CTF/AICD contains an ENTPY consensus sequence that influences internalization in other proteins or can mediate binding to proteins via PTB-conserved motifs that include FE65 (transcriptional control), which also stabilizes AICD, mDAB/abl (neurite arborization), XII/Mints (membrane trafficking), JIP/JNK, numb (Notch signaling), and Shc (signaling) (see below). Among properties proposed for AICD signaling include binding to nuclear transport proteins Mints/XIIs to facilitate the formation of a Fe65histone acetyltransferase/Tip60 complex (Sastre et al., 1998; Biederer et al., 2002; Cao and Sudhof, 2004). RNAi suppression of Mints XIIa/b in transfected neuroglioma cells alters APP degradation (Xie et al., 2005b). PTB domains of nuclear transport proteins constitute drug targets as seen from studies on nuclear receptor corepressor (N-CoR), tetraspanins, Alcadeins, kinases, and munc 18–1 (Baek et al., 2002; Biederer et al., 2002; Ho et al., 2002; King et al., 2004). There is an association between MintsX11s (a, b, g) and Friedrich’s ataxia, and there has been a report for a role for tripartite adherin-like structures of alcadein in APP turnover (Araki et al., 2003). AICD binding to numb or numb-like inhibitors in mouse brain lysates indicates potential cross-talk between APP turnover and Notch signaling pathways that raises concerns for the use of g-SC inhibitors (Roncarati et al., 2002). The function of AICD and its role in AD pathology are the topics of ongoing studies including the creation of Tgs expressing Fe65 alone (Fe.Cr) or in combination with AICD (Ryan and Pimplikar, 2005). AICD Tgs show activation of the Pro-directed Ser/Thr kinase GSK-3b, which phosphorylates tau at multiple sites. The phosphorylation at S9 inhibits GSK-3, but at Y216 activates GSK-3 without evident increase in mRNA by unknown mechanisms. Activation of GSK-3 influences phosphorylation of a number of regulatory and cytoskeletal cell components including MAP1B and cytosolic collapsin responsive mediator protein-1 (CRMP2) that alter microtubule stability and semaphorin 3a/NP-plexin-mediated events on growth cones via GSK-3/CDk pathways (Goold et al., 1999; Cole et al., 2004; Uchida et al., 2005). This has been demonstrated in Tgs although in vivo function remains to be defined in terms of changes in behavior or morphology and relevance to pathology. Among unanswered questions are roles for AICD and turnovers of NFTs, microtubules, axonal growth, and arborization (Leissring et al., 2002; Pardossi-Piquard et al., 2005).

4.5 g-SC Inhibition There is interest in reducing amyloid burden since g-SC is rate limiting for amyloidogenesis (> Figure 16-2). The hypothesis that PS acts as a pseudo-Asp protease led to the synthesis of potent transition-state analogs similar in concept to that for the treatment of AIDS (Shearman et al., 2000; Wolfe, 2001; Tsai et al., 2002). These include pepstatin-like peptidomimetics to suppress APP turnover in transfected cell models (> Table 16-6 for examples and references for L-685,458, LY-411,575, analogs MWIII-36C or -26A, WPE-III-31C, etc.). In TgAPP, daily injections of 0.1–10 mg LY 411575 reduce Abx in CSF fluids, similar to BMS 299897 (Lanz et al., 2004; Barten et al., 2005). LY-411575 being deleterious for the development of peripheral B cells but not T cells illustrates unacceptable side effects arising from impacts on other signaling pathways due to indiscriminate g-SC inhibition. Mechanisms of catalysis are poorly understood, although for a number of years it has been postulated that key PS-Asp groups mediate TM cleavage. This is supported in part by suppression with transition-state analogs or photoaffinity-coupled peptides modeled after those that target viral HIV proteases (Li et al., 2000b; Chun et al., 2004). This has led to the extensive synthesis of derivatives to improve potency, cell penetration, and preference for targeting APP-C99 rather than other TM proteins. These include a large and growing list of difluroketones, aldehydes, expoxides and hydroxy-ethyl, or triamides active at pM-mM levels, but none are yet available for therapy (McLendon et al., 2000; Wolfe, 2001; Bihel et al., 2004; Esler et al., 2004; Prasad et al., 2004; Wrigley et al., 2004). None conform to classical modes of inhibition and exhibit


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. Table 16-6 Inhibitors or strategies to alter g-SC Asp-like site-specific agents Difluoro ketone peptidomimetics (e.g., MW-167 shown, CM-265) (Moore et al., 2000; Wolfe, 2001)

Serine protease-like: JLK2, 6 (Petit et al., 2001)

Proteasome: MG132, lactacystin (Bunnell et al., 1998; Steinhilb et al., 2001) L685,458 (shown) (Shearman et al., 2000); analogs MWlII-36C or -26A

Calpain: z-LL- or VL-cho, N-ac-LLnL-al (Klafki et al., 1996; Figueiredo-Pereira et al., 1999; Verdile et al., 2000) NSAIDs: Eg.: Flurbiprofen

DAPT (Dovey et al., 2001)

Sulindac sulfide

LY-411575 (Lanz et al., 2004) (Eriksen et al., 2003; Weggen et al., 2003; Beher et al., 2004) Benzodiazepines (e.g., compound E shown) (Churcher et al., 2003; Owens et al., 2003) Photoaffinity binding (Esler et al., 2000; Li et al., 2000b)

BMS-289948 (Anderson et al., 2005); MRK-560 active orally 6–10 mg/kg(Best et al., 2006); Fenchylamine sulfonamides (Rishton et al., 2000); (Hydroxyethyl)urea peptides (Wolfe et al., 2002); Pepstatins (Xia et al., 2000; Pinnix et al., 2001); z-IL-cho, boc-K(Dnp)IL-epoxide, z-IL-cho (McLendon et al., 2000); CBAP (Beher et al., 2001)

Gleevec (ST1571) (Netzer et al., 2003) RNAi to silence PS-1/2, aph-1, pen-2, NCT (Francis et al., 2002)

Further examples exist in NCBI databases (http://www.ncbi.nlm.nih.gov/), life science products suppliers (for example, Calbiochem http://www.emdbiosciences.com/; Sigma http://www.sigmaaldrich.com/; Biomol www.biomol.com), or have proprietary designations for clinical evaluation

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nonlinear or noncompetitive kinetics. Furthermore, g-SC is also suppressed by other categories of protease inhibitors or diverse pharmacological agents including sulfonamides, benzodiazepines, Liþ, or GSK inhibitors (Tian et al., 2002; Churcher et al., 2003; Owens et al., 2003; Phiel et al., 2003). A recent study demonstrates that a benzodiazepine derivative, compound E, 30 mg/kg (oral) reduces A40 in guinea pig plasma, CSF, and in the cortex (Grimwood et al., 2005). Combinatorial screening of peptides identified an N-aryl-Ala-ester derivative DAPT and potent benzophenonemethyl analogs as inhibitors of g-SC (Dovey et al., 2001; Kan et al., 2004). Oral administration of DAPT to wild-type Sprague–Dawley rats lowers A40/42 in brain, CSF, and plasma in a dose-dependent manner (El Mouedden et al., 2006). Nonspecific inhibitors generally require higher concentrations and include those found active against proteasome/calpains (MG132, z-LL, VL-CHO, N-ac LLnL) and serine proteases (AESBF, isocoumarins JLK 2–7) (> Table 16-2) (for review see Vassar and Citron, 2000; Esler et al., 2002; Bihel et al., 2003). Allosteric inhibition may contribute to the effects of these agents or of selected NSAIDs (Berezovska et al., 2005; Nakaya et al., 2005). Studies are ongoing to find the possible benefits of long-term administration of the NSAIDs ibuprofen, indomethacin, dapsone, flurbiprofen, sulindac, meclofenamin, and diclofenac or tranquilizer benzodiazepines to delay the onset of dementia (Eriksen et al., 2003; Owens et al., 2003; Sagi et al., 2003; Weggen et al., 2003). It is unclear if NSAIDs act on ancillary pathways that influence a-secretases or Ras-like GTPase Rho signaling (Zhou et al., 2003). Since b-SC and g-SC are rate limiting for amyloidogenesis, drugs that combine inhibition of both enzymes are attractive. Paradoxically, while g-SC mediates normal APP turnover, it is also responsible for the overproduction of Abx that can lead to pathology. Lethality in PS-deficient animals may arise, in part, from the impact of defective turnover of other g-SC substrates such as Notch, which play critical roles in development. Plausibly, a partial suppression of g-SC may have benefits comparable to suboptimal statin dosage by reducing 30–50% the activity of HMG-CoA and amyloidogenesis (Refolo et al., 2001). Equally, differential alteration of rac1, suppression by EHT 1864 (Desire et al., 2005), or use of protein kinase inhibitors to target a g-SC nucleotide-binding site have potential to reduce amyloidogenesis while sparing Notch (Fraering et al., 2005).

4.6 BRI Amyloidoses and lnhibition There is interest in rare British and Danish dementias (FBD-FDD) that yield alternative fibril-forming products unrelated to Ab (> Table 16-2), formed by furin processing of 266-mer precursor proteins. The function of these fibril-forming products are unknown except for their association to an autosomal dominant disorder that shows AD symptoms but with exacerbation of CAA, and in the case of FDD, loss of vision and deafness. Until recently the biochemistry of these disorders was not described although the pathology was published decades ago (Rostagno et al., 2005; see discussions in Online Mendelian Inheritance in Man (OMIM) Internet sites maintained by U.S. NIH (http://www.ncbi.nlm.nih.gov/entrez/ dispomim). The BRI family is an evolutionarily conserved multigene family of precursor proteins (ITM 2A-C, E25 A-C) whose functions are unknown. They contain six exons and five introns with autosomal dominant mutations associated with the long arm of chromosome 13q14 in the case of FBD/FDD (chromosome 13 dementias) with an AD-like pathology. mRNAs are widely distributed in tissues especially in neurons and glia of hippocampus and cerebellum, among others. Structurally this 266-mer protein is a type II glycoprotein (N-terminal intracellular) containing an 100-mer BRICHOS motif in the propeptide region susceptible to furin, a single TM, two cysteines capable of forming a bridge, and one identified glycosylation site at residue 170. Processing at R243-E244 releases fibril-forming peptides with identical 22-mer N-termini but divergent 12-mer C-termini, providing a method to generate distinctive antisera. These cleavages result from a stop codon mutation for FBD or a 10-nt duplication in the case of FDD. Peptides contain N-terminal pyroGlu, which may contribute to the stability of the deposits, but not for circulatory forms. CAA is more pervasive and appears to be associated with members of the complement cascade and furin. Increase in BRI-2 expression reduces Abx whereas RNAi silencing increases b-peptide release, indicating competition for docking with the relevant g-SC sites prior to processing (Matsuda et al., 2005). Recent


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studies show clearance by IDE and that increase in expression of BRI-2 or immunophilin FKB 38 in combination with Mt-bcl-2 reduces Ab amyloidogenesis and CAA. Wang et al., 2005. Also binding of BRI3, a related gene, to BACE-1 shows potential for examining FBD/FDD peptides as lead compounds for mutual interference with APP turnover for attenuating Ab amyloidogenesis (Wickham et al., 2005). There is scope to consider signaling pathways that indirectly alter Ab release such as rac1 and suppression by EHT 1864 (Desire et al., 2005) or protein kinase inhibitors that target a nucleotide-binding site present on g-SC, decreasing Ab secretion while sparing Notch cleavage (Fraering et al., 2005).

5

Obligatory Components of the g-SC

5.1 Presenilins (PS1, PS2) Presenilins coded by chromosome 14 band q.24.3 (PS1, 467-mer, Mr 57 kDa) and chromosome 1 band q.42.1 (PS2, 448-mer) in humans, sharing an 70 homology, were discovered and named so due to their association with FAD (Marks and Berg, 2001). The gene contains ten exons that display tissue-specific alternate splicing. While the function of native nonmutated forms in mammalian tissues remains enigmatic, a number of studies implicate the following roles: Cell viability (Alves da Costa et al., 2002) Binding to receptors/mRNA expression (a-7 nicotinic receptor, Arc, GluR1, mRNAs Nur77 (Dineley et al., 2002; Dickey et al., 2004) Ca2þ capacitance, LTP (Ris et al., 2003) Wnt-catenin signaling pathway (Kang et al., 1999; Uemura et al., 2003) GSK-3/kinesin-I, cell motility in TgPS1M146V cells (Pigino et al., 2003; Sarkar and Das, 2003) A topology comparable to the evolutionarily conserved motif of 7-TM G-protein-coupled receptors (Dewji et al., 2004; Dewji, 2005) Deletion of PS1 is embryonic lethal, indicating a crucial role in development and provides a basis to prepare null (ES) cells for transfection (Feng et al., 2004). The rescue by mammalian PS orthologs of defective egg laying in nematodes and eye/wing development in Drosophila points to roles also in mammalian cells for signaling. To overcome lethality on PS deletion, there are now conditional knockouts of postnatal excitatory neurons, which highlight an association to Ab generation and changes in behavior without altering Notch expression (Yu et al., 2001b). Crossing conditional PS1/ (PS1cKO) with PS2/ mice provides additional models to identify changes in synaptic plasticity in the postnatal forebrain, impairment of memory, and age-dependent neurodegeneration (Saura et al., 2004). This provides a new basis to study correlation to changes in receptors (NMDA), kinases (CREP/CBP), tau (hyperphosphorylation), and amyloidogenesis (Saura et al., 2005). Hydropathic plots for PS indicate ten hydrophobic regions (HR) almost identical to those of the nematode homolog sel-12 indicating that secondary structure contributes to function. This is consistent with 7–9 TM although subject to dispute (Lehmann et al., 1997; Li and Greenwald, 1998; Nakai et al., 1999; Dewji et al., 2004). For example, sel-12 constructs with b-galactosidase insertion after HR-2, -4, -6, - 7, -9 and -10 favor an 8-TM model with the key Asp groups thought to be responsible for catalysis on TM–6 and-7 (Li and Greenwald, 1998). Surface epitope labeling on PS1 sites yields variable data that differ in orientation of Asp groups (Dewji et al., 2004; Brunkan et al., 2005; Laudon et al., 2005; Oh and Turner, 2005), leading to questions on the number necessary for catalysis (Sisodia et al., 2001). There is also interest in TMs that mediate PS endoproteolysis to form active fragments (Brunkan et al., 2005), and whether FAD mutations significantly affect binding to g-SC and hence specificity toward APP/C-99 cleavage or other proteins (Walker et al., 2005). New studies identify a number of alternative proteins capable of binding to PS that modulate Ab production and trafficking via mechanisms that remain to be identified. These include ubiquilin to influence PS endoproteolysis and alter pen-2 interaction with NCT (Massey et al., 2005), and phospholipase D1 that alters APP trafficking and neurite outgrowth, implying an association with lipid metabolism (Cai et al., 2006).

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The fact that PSDex9 retains toxicity has not fully been explained since it is not subject to endoproteolysis (Nakaya et al., 2005; Kim et al., 2005b). One conclusion is that the release of 27–28 kDa N-terminal and 18–20 kDa C-terminal metabolites (Met292 or Met298 cleavage for PS-1 or -2 respectively; > Figure 16-9) is not mandatory for the successful formation of an active g-SC complex. Endoproteolysis has attracted

. Figure 16-9 Topological model for PS-1. (a) Hydrophobic regions (HR) of PS-1 and the Caenorhabditis elegans homolog sel12. (b) Postulated topology of PS-1 with 7-TM ( ) and remaining extracellular and intracellular HR domains ( ) (Dewji et al., 2004). Key Asp residues 257 (HR 6) and 385 (HR 8) shown ( ). Major and minor sites of PS-1/2 endoproteolysis (—>,--->) releasing major NTF/CTFs are shown in inset ( ). FAD point mutations for PS-1 (○) and PS-2 (●) and sequence deletion for the biostable Dexon9 ( ) shown

attention as one method to modulate g-SC activity assuming this is essential for the majority of PS mutants. Suppression by CM35 or MW167 suggests an unknown aspartyl protease (Campbell et al., 2003; Xia, 2003). Suppression by a peptide encompassing the susceptible loop region (IC50 ¼ 2.1 mM) acting via an L684,458-insensitive pathway (Knappenberger et al., 2004) can supply a lead compound (Dewji, 2005). Early studies focused on PS sequences critical for ‘‘toxicity’’ and were conducted before the realization of the existence of a multimeric complex. Progress has been slow but suggests critical roles for a CTF 349–467 domain (PAL or P414ALP) and/or the NTF (Tyr288) (Tomita et al., 2001; Kim et al., 2005b; Wang et al., 2006a). Chimeric PS, composed of NTF from PS1 and CTF from PS2, retain activity within g-SC and are labile, forming separate fragments on endoproteolysis. Transfection of NTF and CTF fragments of differing origin reconstitute g-SC, indicating conservation of critical consensus sequences among different species (Stromberg et al., 2005). Interestingly, PS1 NTF and a caspase-generated CTF (C31) can also form an active g-SC when coexpressed in SH-SY5Y cells deficient in PS (Hansson et al., 2006). Deletion of the


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hydrophilic loop (exon 10) reduces A40 but not A42 pathology in Tgs, suggesting A40 is protective in this mutant (Deng et al., 2006). One of the most controversial areas is the inference that PS acts as an actual cryptic Asp-like protease despite lacking a classical enzymatic center. It is possible this is the case, but within the new context of a configuration within a multimeric complex. Several groups point to similarities with several bacterial signal peptide peptidases (SPP) containing a Gly-X-Gly-Asp385-X motif or polytopic type 4 prepilins (Steiner et al., 2000; Moliaka et al., 2004; Nyborg et al., 2004). Swapping this motif to create a chimera with a distant homologous peptide SPE-4 supports APP but not Notch processing, indicating a role in determining specificity (Yamasaki et al., 2006). Interestingly, the C-terminal PAL motif of PS1 noted above is also conserved in SPP (Wang et al., 2006a). One member of this family, SPPL2, is localized in endosomal– lysosomal vesicles (Krawitz et al., 2005). An important issue is how multimeric complexes act on peptide bonds buried as a-helices in a hydrophobic environment protected by hydrogen bonding. Arguably these may form substructures to insulate them from lipid bilayers, similar to those postulated for aquaporins and the KcsA Kþ channel proteins (Hamasaki et al., 2002). Similarly g-SC is a membrane component, which presents new challenges for its isolation, when extracting, without disturbing its intimate association with the substrate.

5.2 Nicastrin Isolation of NCT from immunoextracts of HEK293PS1 cells and its binding to PS1 provided impetus to identify multimeric complexes for RIP (Yu et al., 2000). NCT, a 709-mer protein homologous to C. elegans aph-2, is evolutionarily conserved and is necessary to facilitate Notch processing/signaling (Goutte et al., 2000; Edbauer et al., 2002b; Kaether et al., 2002; Herreman et al., 2003; Morais et al., 2003; Nyabi et al., 2003). It is ubiquitously distributed in tissues, also present in neurons as a spliced form lacking the 62-mer coded by exon 3 (Ilaya et al., 2004; Confaloni et al., 2005). It associates also with cerebellar synapses during development (Uchihara et al., 2006). The deduced sequence has a single TM, a D336YIGS motif, multiple sites for glycosylation capable of providing a scaffold for the assembly of other g-SC components and targeting to the appropriate compartments (Hattori et al., 2002; Tomita et al., 2002; Capell et al., 2003; Morais et al., 2003; Shirotani et al., 2003; Kaether et al., 2004; Kim et al., 2004). Deletion of DYIGS reduces A42/40 secretion with accumulation of C83/C99 rather than continued processing to liberate Abx (Yu et al., 2000). Interaction of the TM of NCT with the C terminus of PS is proposed as a potential mechanism to facilitate assembly and activation of g-SC (Morais et al., 2003; Kaether et al., 2004). Evidently NCT is critical for development since its deletion is embryonic lethal at day 10.5, yielding deficient null cells similar to PS1/2/ (Li et al., 2003b). This is a useful feature since homozygous fibroblasts from NCT nulls are defective in APP processing when compared to heterozygotes, which show a gene-related 50% decrease in Abx secretion. In addition, null fibroblasts contain lower amounts of aph-1 and pen-2, in line with lower amounts of PS fragments (Zhang et al., 2005b). A point mutation in the PS ectodomain abolishes NCT maturation and impairs g-SC processing of APP and Notch (Olry et al., 2005) and conforms to the view that both components are essential for processing. Furthermore, this is supported by NCT RNAi, which results in a decrease in APP catabolism (Edbauer et al., 2002b).

5.3 Aph-1 This 7-TM protein, a highly conserved protein in different species (mammals, Danio, Drosophila, and Arabidopsis), is essential for Notch processing and signaling (Ma et al., 2005; Niimura et al., 2005). Alternative splicing results either in a fragment with a C-terminal KD or a longer one with RRQEDSRVMVYSALRIPPED. Depending on the species, there also are isoforms aph 1a (in humans coded by chromosome 1)(Ma et al., 2005), aph-1b (chromosome 15), 1c (rodents) (Serneels et al., 2005), or an aph-1b that lacks TM-4 coded by exon 4 (Saito et al., 2005a).

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Studies on nematodes establish a critical role for G122XXXG on TM-4 of aph 1a in development or for TM cleavage on transfection of mutant form lacking this motif in K293 cells (Edbauer et al., 2004), indicating it as essential for assembly and maturation of multimeric complexes (Lee et al., 2004; Hansson et al., 2005). Aph-1a deletion is embryonic lethal, demonstrating an essential role in development (Serneels et al., 2005). Few pharmacological studies are currently available except that a lower expression of aph-1b is seen in apomorphine-susceptible rats (Coolen et al., 2005).

5.4 Pen-2 Pen-2 (presenilin enhancer 2), a 2-TM 101-mer protein (human chromosome 19q13.1), lacks a signal peptide and was discovered in nematode DNA screens as one of the components essential for development via Notch processing and signaling (> Table 16-1). Little has been established on transcriptional control, other than the presence of a 238-bp sequence located 353 bp upstream of the ORF, containing a CREBbinding domain. Activation of CREB by forskolin promotes pen-2 without altering other obligatory components (Wang et al., 2006b). Its wide distribution in species (similar to those for aph-1) suggests that these proteins coordinate to trigger the catalytic cascade within a multimeric matrix (Francis et al., 2002). The use of pen-2 constructs implicates essential roles for C-terminal D90YLSF and for one or more N-terminal motifs to bind PS (Crystal et al., 2003; Hasegawa et al., 2004; Prokop et al., 2004) and also for the proximal two-thirds of TM-1 in the cleavage of labile forms of PS-1 (Kim and Sisodia, 2005a). The cytosolic loop between the two TMs protects Danio embryos from caspase-dependent apoptosis: this implies a role for this motif in development (Zetterberg et al., 2006). Haplotype-based analyses do not implicate pen-2 to be a significant risk factor in FAD (Bertram et al., 2004). New studies suggest PS-TM4 interacts with pen-2 via conserved motif W203NF or N204F to facilitate endoproteolysis and also with g-SC (Kim and Sisodia, 2005b; Watanabe et al., 2005).

6

Alternative Proteases as Secretases

APP, as a 100–120-kDa glycosylated isoform, contains peptide sites vulnerable to other proteases. Depending on localization and the environment (pH, cofactors, inhibitors), these may act to process APP or permit cross-talk with other pathways for processing (Nixon et al., 2001). While Abx predominates in SPs, other components (cathepsins, metalloproteases, proteoglycans, complement, metals) (Marks and Berg, 1997) ostensibly play supplementary roles in APP catabolism. Cathepsins B, D, and S, due to their specificity, appear unlikely to act directly as bonafide secretases, although they may play supplementary roles in processing APP in secretory vesicles, based on a study in chromaffin cells using the inhibitor CA 074 methyl ester (Hook et al., 2005). Cathepsin D polymorphisms which are potential risk factors associated with AD do not alter Abx release, as seen from a study on KOs and have no direct linkage in an established AD Spanish cohort (Papassotiropoulos et al., 1999; Mateo et al., 2002). But recently, an Ala220-Val224 shift within exon 2 has been reported to modulate ApoE effects on fibrillogenesis (Davidson et al., 2006). Lysosomal cathepsins B and L degrade A40 at F19-F20 suggesting roles in its clearance but not necessarily in its formation (Marks et al., 1994), although in a study on chromaffin granules an inhibitor CA 074Me reduces APP processing (Hook et al., 2005). Caspases are cell death Asp-specific proteases that increase in apoptosis in vitro or in brain trauma resulting from ischemia, stroke, or radiation (Gervais et al., 1999; Marks and Berg, 1999). Like cathepsins they lack specificity in order to classify them as secretases per se, although they can degrade APP at alternative sites (Loetscher et al., 1997). Cleavage of C-99 generates C-31, which decreases viability of cells expressing APP via a pathway sensitive to caspase inhibitors (Lu et al., 2000). Inhibitors of other cell death modulators of the caspase pathways attenuate Abx toxicity in vitro mediated by cytosolic caspases, or in the case of caspase-12 by gene deletion of an ER protein responding to Abx toxicity (Nakagawa et al., 2000).


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Conclusion

The use of APP as a prototypic type-1 substrate has provided new insights in the enzymology of converting enzymes and protein turnover. First, it led to the discovery of a novel family of Asp proteases (BACE). Second, it established a new role for metalloproteases of the ADAM family as putative a-secretases. Third, it provided the basis for identifying multimeric complexes to mediate the final release of Abx. Among unresolved questions are whether there is a specific a-secretase, what is the function of intracellular APP, and how mutations in it or other FAD proteins confer a genetic autosomal dominant gain-of-function in terms of amyloidogenesis. Cleavage of multiple substrates by b- and g-secretases presents issues for design of site-specific agents to target amyloidogenesis without incurring collateral damage to other cellular events or associated signaling pathways. Ultimately, as is the case for other CNS disorders, there is a need to design agents capable of penetrating to cellular sites relevant to APP turnover. Precedents have been established to use protease inhibitors to treat hypertension or AIDS, which augurs well to consider peptides or peptidomimetrics for this goal. Alternatively this can supplement other initiatives involving b-pleated disruptive agents, immunotherapy, or activation of a-secretase to promote the nonamyloidogenic pathway. The postulate that key Asp groups of PS form a catalytic center has led to synthesis of potent transitionstate inhibitors, but these do not explain effects of other categories of protease inhibitors or action of antianxiolytic benzodiazepines, antiinflammatory NSAIDs, and cholesterol-lowering statins which also attenuate amyloidogenesis. Many of these appear to interact with allosteric sites of g-SC, but a full explanation is awaited. Future progress will depend on isolation of homogenous g-SC to reveal the alignment of components, mechanisms for intramolecular PS proteolysis, docking with substrate, and subsequent cleavage within the hydrophobic environment of the TM. Application of mass spectrometry and proteomics can provide new insights into topology and mechanisms of assembly of obligatory components including stable and labile forms of PS1/2 or their fragments. As such the study of secretases opens a new chapter in the enzymology of protein turnover that can provide hope to develop therapies to attenuate recalcitrant forms of AD dementia.

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The Biology of Caspases in Central Nervous System Trauma

M. L. McEwen . J. E. Springer

1

Apoptosis Versus Programmed Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 516

2 2.1 2.2 2.3

The Structure and Function of Caspases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 517 Caspase Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 517 Inflammatory Caspases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 517 Caspases of the Death Cascade . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 519

3 3.1 3.2 3.3 3.4 3.5

Pathways of Apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 520 The Mitochondrial (Intrinsic) Pathway of Apoptotic Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 520 Regulation of Apoptosis by Bcl‐2 Family Members at Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 521 The Death Receptor (Extrinsic) Pathway of Apoptotic Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 525 The ER (Intrinsic) Pathway of Apoptotic Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 526 Additional Pathways of Caspase Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 528

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Substrate Cleavage by Activated Caspases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 529

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Caspase‐Independent Apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 530

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Non‐Apoptotic Functions of Caspases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 530

7 7.1 7.2 7.3 7.4

The Role of Caspases in Neurotrauma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 530 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 530 Ischemic Brain Injury (Stroke) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 531 Traumatic Brain Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 531 Traumatic Spinal Cord Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 532

8 8.1 8.2 8.3

Therapeutic Interventions in CNS Trauma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 533 Competitive Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 533 Anti‐Apoptotic Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 534 Targeting the Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 535

9

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 535

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The biology of caspases in central nervous system trauma

Abstract: Programmed cell death (PCD) is a naturally occurring event in the developing organism, whereby damaged, mislocated, or superfluous cells are discretely and efficiently removed. All of our cells contain the machinery required for their own demise, which occurs through a process of apoptosis and must be highly regulated. Recent research has shown that the apoptotic machinery in many cell types is activated in the central nervous system (CNS) following a variety of insults. Once apoptosis is initiated, an intricate process ensures activation of a family of cysteine proteases called caspases, which are responsible for many of the morphological and biochemical features of apoptosis. Currently, 14 caspases have been identified. In this chapter, we begin with a brief discussion of the structure and function of the caspases. Next, several pathways of apoptotic cell death will be presented, focusing on caspase participation. The chapter concludes with a discussion of data regarding caspase activation following CNS insults, as well as potential therapeutic strategies that target various events in the apoptotic process.

1

Apoptosis Versus Programmed Cell Death

Cell death is a naturally occurring event that eliminates damaged, mislocated, or superfluous cells during the development of multicellular organisms, and regulates tissue homeostasis in adults (Glucksmann, 1951; Saunders, 1966; Wyllie et al., 1980; Oppenheim, 1991; Raff et al., 1993; Vaux and Korsmeyer, 1999; Buss and Oppenheim, 2004). Deregulation of cell death can result in pathological conditions that range from cancer (a paucity of cell death; Strasser et al., 1990) to autoimmune disease (enhanced cell death; Strasser et al., 1991). Morphological forms of cell death have roughly been classified as necrotic or apoptotic. Necrosis is the cell death process by which a cell explodes and spills its intracellular contents. Characteristics of necrotic cell death include disruption of the plasma membrane, cytoplasmic and mitochondrial swelling, random DNA cleavage, and the induction of a pronounced inflammatory response. Apoptosis is a tidy process whereby cells commit suicide and reduce themselves to tiny packages that are devoured by neighboring phagocytes. Characteristics of apoptotic cell death include cytoplasmic shrinking, chromatin condensation, membrane blebbing, fragmentation of intranucleosomal DNA into 200‐bp segments, and the formation of apoptotic bodies with minimal inflammatory response (Kerr et al., 1972). ‘‘Apoptosis’’ was introduced to define a particular type of cell death with specific morphological characteristics. ‘‘Programmed cell death’’ (PCD) is the series of coordinated, genetically controlled events that lead to cellular suicide regardless of the morphological features (Schwartz and Osborne, 1993). Because cells undergoing PCD often have an apoptotic morphology, PCD and apoptosis are usually equated. However, the terms are not synonymous (Assuncao Guimaraes and Linden, 2004). Cells undergoing PCD in vertebrate and nonvertebrate systems do not always display an apoptotic morphology (Schweichel and Merker, 1973; Clarke, 1990; Schwartz et al., 1993; Kitanaka and Kuchino, 1999; Leist and Jaattela, 2001). Terms such as oncosis (Majno and Joris, 1995), autophagic cell death (Clarke, 1990; Bursch, 2001, 2004; Lockshin and Zakeri, 2004), or parapoptosis (Sperandio et al., 2000) describe cells undergoing PCD with morphologies that vary from classical apoptosis or classical necrosis. The primary classifications of cell death (necrosis and apoptosis) probably do not represent two distinct modes of cell death, but the extremes on a cell death continuum (Portera‐Cailliau et al., 1997a, b; Zeiss, 2003). If necrosis and apoptosis were discrete forms of death, one would expect that each form of death would be initiated by different stimuli and that the underlying mechanisms to execute the demise of the cell would differ. However, the same stimulus can initiate apoptosis or necrosis, depending on the mitochondrial response (Ankarcrona et al., 1995; Leist et al., 1997, 1999), and inhibition of one form of cell death does not always lead to cell survival, but may switch the form of the death (Hirsch et al., 1997; Leist et al., 1997, 1999; Kitanaka and Kuchino, 1999). Because of its relative simplicity, the nematode, Caenorhabditis elegans, has been widely used as a model to understand the molecular mechanisms responsible for PCD. The same 131 of 1090 neurons die by PCD during the development of this organism, so this system can be used to map the genes responsible for particular cell fates (Sulston, 1976). Ellis and Horvitz (1986) determined that the actions of two gene products, Ced‐3 and Ced‐4, were largely responsible for all cell deaths during the development of this worm.


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Another gene, Ced‐9, was required to inhibit PCD (Hengartner et al., 1992). In contrast, the cell death machinery has become much more complex in vertebrates (Aravind et al., 2001). Not only are there additional ‘‘players’’ that are not found in C. elegans (e.g., cytochrome c; also known as apoptotic protease‐ activating factor‐2 (Apaf‐2); Putcha and Johnson, 2004), but single gene products with roles in cell death or survival in the nematode may be homologous to ‘‘families’’ of proteins in mammals. The Ced‐4 gene product is homologous to the mammalian Apaf‐1 (Zou et al., 1997), but the Ced‐3 gene product is homologous to the mammalian family of proteases, called caspases (Yuan et al., 1993), and the Ced‐9 gene product is homologous to the mammalian family of Bcl‐2‐like proteins (Hengartner and Horvitz, 1994). Cps‐6, which is homologous to the mammalian endonuclease G (Endo G) (Parrish et al., 2001), and Wah‐1, which is homologous to the mammalian apoptosis‐inducing factor (AIF) (Wang et al., 2002) were the first mitochondrial proteins identified to participate in apoptosis in C. elegans (Parrish et al., 2001). In contrast, several mitochondrial proapoptotic factors have been identified in mammals (cytochrome c, Smac/Diablo, AIF, Endo G, HtrA2/Omi). Caspases (cysteine aspartic acid‐specific proteases) are activated in most cases of PCD with apoptotic morphology, which has prompted suggestions to redefine apoptosis as cell death that requires caspase activation for its characteristic morphological features (Samali et al., 1999; Putcha and Johnson, 2004). However, there are cases where caspases are not activated and cells die with an apoptosis‐like morphology (Adjei et al., 1996; Miller et al., 1997; Lavoie et al., 1998; Susin et al., 1999b, 2000; Doonan et al., 2003), as well as cases where caspases are activated in cells dying with a necrosis‐like morphology (Shimizu et al., 1996; Sperandio et al., 2000). Although the latter may represent cells that initially began dying by apoptosis but switched to necrosis, accumulating evidence suggests that necrosis‐like cell death may also require the intracellular signaling pathways thought to direct apoptosis (Kawahara et al., 1998; Kitanaka and Kuchino, 1999). Clearly, cell death is a complicated process. The form of cell death and the underlying mechanisms are probably dependent on numerous factors, such as cell age, cell type, death stimulus, and environmental milieu. The purpose of this review is not to discuss all forms of cell death, but to characterize the mammalian family of caspases and illuminate their roles in PCD following injury to the mature CNS.

2

The Structure and Function of Caspases

2.1 Caspase Structure The caspases are an evolutionarily conserved family of proapoptotic cysteine‐dependent proteases that play an integral role in cell death in all metazoans. Caspases are present in all healthy cells as inactive proenzymes (zymogens) that are approximately 30–50 kDa. Caspases have (1) an N‐terminal prodomain that varies in length from 22 to more than 200 amino acids, (2) a large subunit approximately 17–20 kDa, (3) an intersubunit region of approximately 10 amino acids, and (4) a small subunit approximately 10–12 kDa (> Figure 17-1). Caspase activation requires that the procaspases be cleaved into their large and small subunits, which then form heterotetramers from the association of two heterodimers. Each caspase heterodimer consists of a large and small caspase subunit (Cohen, 1997; Donepudi and Grutter, 2002; Shi, 2002). Currently, 14 members of the caspase family have been identified in mammals and can be categorized by their structure and function. Caspases have roughly been categorized as mediators of inflammation, initiators of the death cascade, or executioners of the death cascade. Twelve of the 14 mammalian caspases have generally been identified in humans, rodents, and other mammals, whereas caspases‐11 and ‐12 have only clearly been identified in rodents (Alnemri et al., 1996; Hu et al., 1998a; Thornberry and Lazebnik, 1998; Van de Craen et al., 1998; Wang et al., 1998; Lamkanfi et al., 2002).

2.2 Inflammatory Caspases Caspase‐1, originally named interleukin‐1b‐converting enzyme (ICE), caspase‐4 (ICErel‐II, TX, ICH‐2), caspase‐5 (ICErel‐III, TY), and caspase‐11 are the inflammatory caspases, which have only been identified in

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. Figure 17-1 This diagram illustrates the general structure of procaspases‐1 through ‐12. The inflammatory caspases and the initiator caspases have a large prodomain that contains either a caspase‐recruiting domain (CARD) or two death effector domains (DED) to mediate protein–protein interactions. Executioner caspases have very short prodomains, if any

vertebrates (Lamkanfi et al., 2002). Although there is some evidence that these caspases participate in apoptosis (Fink et al., 1999; Schielke et al., 1998; Li et al., 2000; Rabuffetti et al., 2000), they are primarily involved in the processing of proinflammatory cytokines, such as IL‐1b and IL‐18 (Martinon and Tschopp, 2004). The inflammatory caspases have a long, N‐terminal prodomain that contains an 80–90‐amino‐acid motif called a caspase‐recruiting domain (CARD) (see > Figure 17-1), which mediates signaling through homophilic protein–protein interactions (Hofmann, 1999). However, the pathways that activate the proinflammatory caspases are not well studied. In general, their activation appears to require recruitment to a caspase activation platform, which brings the caspases into close proximity to one another and causes them to autoactivate (Martinon and Tschopp, 2004). When tumor necrosis factor (TNF) binds to the TNF death receptor (TNFR), the receptors cluster at the plasma membrane and cause clustering of the receptors’ intracellular death domains (DDs). DDs are regions of approximately 80 residues at the C‐terminal that are required for homophilic protein–protein interactions (Hofmann, 1999). The TNFR‐associated death domain (TRADD) is then recruited to the receptor aggregates via its DD to serve as an intermediate adapter to recruit the DD‐containing proteins TNFR‐associated factor‐2 (TRAF‐2), receptor‐interacting protein (RIP), and cIAP1, which is a member of the inhibitors of apoptosis (IAP) family (see > Figure 17-5). Together, this complex causes activation of the proinflammatory pathway through NF‐kB and jun N‐ terminal kinase signaling. Activated NF‐kB leads to expression of cFLIP (FLICE (Fas‐like interleukin‐1b‐converting enzyme)‐like inhibitor protein), which inhibits caspase‐8 activation at the receptor complex and inhibits the death receptor‐mediated (extrinsic) pathway of apoptosis (Danial and Korsmeyer, 2004). The CARD‐containing ICE‐associated kinase (CARDIAK; also known as RIP2, Rick, CCK2, CARD3) is a serine/threonine kinase that can bind with the DD of TRAF‐2 and the CARD of procaspase‐1 suggesting that CARDIAK is an adapter molecule that recruits caspase‐1 to the signaling cascade of the TNFR family members (Thome et al., 1998). Caspase‐11 physically interacts with procaspase‐1 and may be required for caspase‐1 activation


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(Wang et al., 1998). Several other CARD‐containing proteins have been identified in recent years, which interact with CARDIAK and/or caspase‐1 to manipulate activation of proinflammatory pathways. Because the focus of this chapter is on the role of caspases in cell death, the inflammatory pathways will not be discussed further.

2.3 Caspases of the Death Cascade Participants in the death cascade have been subdivided into two groups based on the length of their prodomain and its associated function. Class I procaspases are generally monomers (except procaspase‐2) that have relatively long prodomains, which enables oligomerization of the procaspase molecules. Because class I caspases can dimerize and autoactivate (Boatright et al., 2003; Donepudi et al., 2003), they are also known as initiator caspases and include caspases‐2 (ICH‐1), ‐8 (FLICE, MACH, Mch5), ‐9 (ICE‐LAP6, Mch6, Apaf‐3), and ‐10 (Mch4). Although sometimes classified as an inflammatory caspase by its structural similarity to the other inflammatory caspases (Van de Craen et al., 1997; Lamkanfi et al., 2004), caspase‐12 has been classified as an initiator caspase because of its function in endoplasmic reticulum (ER) stress‐ induced apoptosis (Nakagawa and Yuan, 2000; Nakagawa et al., 2000; Rao et al., 2001, 2002a; Yoneda et al., 2001; Szegezdi et al., 2003). This classification is used in this chapter. All class I procaspases have N‐terminal regions of complementary binding capability that mediate protein–protein interactions (see > Figure 17-1). Procaspases‐2, ‐9, and ‐12 have only one of these regions, called a CARD, whereas procaspases‐8 and ‐10 have two regions, known as death effector domains (DEDs; Hofmann, 1999). These N‐terminal domains are removed and the procaspases are cleaved within their intersubunit region to create the processed (active) caspase. Class II procaspases are dimers that have short prodomains and are activated by limited proteolysis within their intersubunit region (Kumar and Colussi, 1999; Chai et al., 2001; Riedl et al., 2001; Boatright et al., 2003). Because they lack the long prodomain (see > Figure 17-1), class II procaspases must rely on initiator caspases for cleavage and activation. Therefore, class II caspases, which includes caspase‐3 (CPP32, apopain, Yama), ‐6 (Mch2), and ‐7 (Mch3, ICE‐LAP3, CMH‐1), are known as executioner caspases (Kumar, 1999). The actions of these caspases are primarily responsible for the morphological features of apoptosis. Caspase‐13 (also known as evolutionarily related ICE (ERICE)) shares sequence homology with the inflammatory caspases. Unlike other members of the ICE subfamily, caspase‐13 is activated by caspase‐8, which suggests a downstream role for caspase‐13 in apoptosis (Humke et al., 1998). More recent evidence has suggested that caspase‐13 is not actually a human caspase (Koenig et al., 2001), which may explain why little else is known about its role in apoptosis. Caspase‐14 (also known as mini‐ICE (MICE)) also shares sequence homology with the inflammatory caspases. However, procaspase‐14 is a monomer that lacks an N‐terminal prodomain and CARD (Ahmad et al., 1998; Hu et al., 1998a). Similar to other caspases, procaspase‐14 requires cleavage between the large and small subunits and subsequent dimerization for activation (Mikolajczyk et al., 2004). Caspase‐14 may be activated following trauma (Krajewska et al., 2004) and during death receptor‐ or granzyme B‐mediated apoptosis (Ahmad et al., 1998; but see Hu et al., 1998a) suggesting that caspase‐14 has a downstream role in cell death. However, procaspase‐14 is only weakly processed by caspase‐8 or ‐10, and may require cleavage by granzyme B or calpain (Ahmad et al., 1998; Van de Craen et al., 1998; Hu et al., 1998a; Krajewska et al., 2004; Mikolajczyk et al., 2004). However, the functional characteristics of caspase‐14 vary by species (Mikolajczyk et al., 2004). The natural substrates of caspase‐14 are yet to be identified, but the abundance of caspase‐14 in embryonic tissues and in adult skin have suggested a primary role for caspase‐14 in ontogenesis and adult skin physiology (Van de Craen et al., 1998; Hu et al., 1998a; Chien et al., 2002; Alibardi et al., 2004; Walsh et al., 2005). With the exceptions of caspases‐13 and ‐14, activated caspases cleave and activate other caspases to amplify the cascade of caspase activation and cellular destruction. The two most thoroughly described pathways of caspase‐dependent cell death are the mitochondria‐initiated (intrinsic) pathway and the cell‐ surface death receptor (extrinsic) pathway. Recent evidence suggests that the ER serves as a third cellular compartment (intrinsic) that regulates apoptosis.

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Pathways of Apoptosis

3.1 The Mitochondrial (Intrinsic) Pathway of Apoptotic Cell Death Several proteins important for mitochondrial homeostasis under nonapoptotic conditions participate in apoptotic cell death (Reed, 1997; Daugas et al., 2000a; Lipton and Bossy‐Wetzel, 2002; Lorenzo and Susin, 2004; Vahsen et al., 2004). The intrinsic apoptotic pathway requires that unknown intracellular stimuli cause the release of these proapoptotic factors from mitochondria into the cytoplasm. Released cytochrome c (Kluck et al., 1997; Yang et al., 1997; Finucane et al., 1999) interacts with Apaf‐1 to form a caspase recruitment platform, called the apoptosome (Adams and Cory, 2002; Hill et al., 2003). Apaf‐1, which has an N‐terminal CARD, is inert in the cytoplasm until it interacts with cytochrome c in the presence of adenine triphosphate (ATP). Procaspase‐9 is then recruited to the apoptosome via its N‐terminal CARD (> Figure 17-2). When a sufficient number of caspase‐9 zymogens are in close proximity, caspase‐9 autoactivates (Liu et al., 1996, Li et al., 1997b; Srinivasula et al., 1998; Hu et al., 1998b, 1999; Salvesen and Dixit, 1999; Zou et al., 1999), which is important for subsequent activation of downstream executioner caspases (Li et al., 1997b; Hu et al., 1999; Slee et al., 1999; Hill et al., 2003). Some evidence suggests that procaspase‐9 recruits procaspase‐3 to the apoptosome for activation (Hu et al., 1999; Bratton et al., 2001). Activated caspase‐3 then cleaves procaspases‐2 and ‐6, and activated caspase‐6 activates caspases‐8 and ‐10. Activated caspase‐3 can also feedback to propagate further activation of caspase‐9 (Slee et al., 1999; Hill et al., 2003). In addition to activating one another, activated caspases cleave a variety of cellular proteins that cause the morphological features of apoptosis (e.g., cytoplasmic shrinking, membrane blebbing, DNA fragmentation), and cleave proteins important for repair. Permeabilized mitochondria also release Smac/Diablo (Du et al., 2000; Verhagen et al., 2000; Adrain et al., 2001) and HtrA2/Omi (Hegde et al., 2002; Martins et al., 2002; van Loo et al., 2002; Verhagen et al., 2002) during apoptosis. These proapoptotic factors facilitate caspase activation by inhibiting the antiapoptotic activities of members of the mammalian IAP family (> Figure 17-2). The IAPs normally inhibit apoptosis by binding activated initiator and executioner caspases and preventing downstream events or by interfering with the processing of the procaspases (Deveraux et al., 1997, 1998; Roy et al., 1997; Salvesen and Duckett, 2002). Smac/Diablo (Srinivasula et al., 2000, 2001; Verhagen et al., 2000; Wu et al., 2000) and HtrA2/Omi (Suzuki et al., 2001; Hegde et al., 2002; van Loo et al., 2002; Verhagen et al., 2002) inhibit the IAPs by physically binding to the IAPs and preventing their interaction with caspases. HtrA2/Omi may also cleave IAPs through its serine protease activity (Srinivasula et al., 2003; Yang et al., 2003). In addition, mitochondria release AIF and Endo G (> Figure 17-2), which translocate to the nucleus and cause chromatin condensation and large‐scale DNA fragmentation (Susin et al., 1999b; Daugas et al., 2000a; Li et al., 2001; Loeffler et al., 2001; van Loo et al., 2001; Cande et al., 2002; Cregan et al., 2002, 2004; Yu et al., 2002). Direct binding between AIF and DNA is important for DNA cleavage (Ye et al., 2002), but AIF cannot cleave DNA (Susin et al., 1999b, 2000). AIF can cause activation of caspase‐9 (Susin et al., 1999b), which could lead to caspase‐dependent cleavage of the inhibitor of caspase‐activated DNase (ICAD). Once free from its inhibitor, caspase‐activated DNase (CAD) could translocate to the nucleus to cleave DNA (Enari et al., 1998). In the absence of caspase activity, granzyme B can cleave ICAD (Thomas et al., 2000; Sharif‐Askari et al., 2001) in the same cleavage site as activated caspase‐3 (Sharif‐Askari et al., 2001) to free CAD. However, CAD may not be the only endonuclease capable of actually cleaving genomic DNA (McIlroy et al., 2000; Yakovlev et al., 2001). One candidate is Endo G (Li et al., 2001; van Loo et al., 2001), which normally participates in the replication of mitochondrial DNA (Cote and Ruiz‐Carrillo, 1993). It is interesting to note that mitochondrial pools of procaspases‐2, ‐3, and ‐9 have been identified and may be released into the cytoplasm during apoptosis (Mancini et al., 1998; Zhivotovsky et al., 1999; Susin et al., 1999a; Costantini et al., 2002). Mitochondrial procaspase‐3 was even colocalized with cytochrome c (Mancini et al., 1998). Currently, the mechanisms responsible for releasing these procaspases are not known, or if mitochondrial release into the cytoplasm is required for their activation.


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. Figure 17-2 This diagram highlights some of the participants in the mitochondrial (intrinsic) apoptotic pathway. A black ‘‘X’’ indicates a protein or event that is inhibited by the preceding component

3.2 Regulation of Apoptosis by Bcl‐2 Family Members at Mitochondria Mitochondrial detection of an apoptotic stimulus initiates a cascade of events whereby mitochondria release factors into the cytoplasm, causing an amplifying cascade of caspase activation, destruction of

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cellular proteins, destruction of DNA, inhibition of antiapoptotic mechanisms, and inhibition of cellular repair. Therefore, it is easy to understand why the release of proapoptotic factors, such as cytochrome c, from mitochondria has been considered the step that commits a cell to apoptotic death (Green and Kroemer, 1998) and that sophisticated mechanisms are necessary to regulate apoptotic pathways. The mechanisms responsible for initiating the mitochondrial pathway of apoptosis are not completely understood but are dependent on a complicated interplay and balance between anti‐ and proapoptotic members of the Bcl‐2 family (Korsmeyer et al., 1993; Yang et al., 1997; Adams and Cory, 1998, 2001; Finucane et al., 1999; Cheng et al., 2001; Cory and Adams, 2002; Scorrano et al., 2003). Several Bcl‐2 family members contain a C‐terminal ‘‘anchor sequence’’ (> Figure 17-3), which targets them to membranes of the mitochondria, ER, and the nuclear envelope (Hockenbery et al., 1990; Krajewski et al., 1993; Ng et al., 1997; Hacki et al., 2000; Germain et al., 2002; Rudner et al., 2002; Mund et al., 2003; Scorrano et al., 2003; Thomenius et al., 2003; Zong et al., 2003; Annis et al., 2004). However, the conformational structure of . Figure 17-3 This diagram depicts the structure of representative Bcl‐2 family members. Subdivisions within this family are based on the type and number of Bcl‐2 homology (BH) domains that the family members share. In general, Bcl‐ 2‐like antiapoptotic family members share sequence homology in all four domains. Bax‐like proapoptotic family members generally share sequence homology in regions 1 to 3, whereas BH3‐only proapoptotic family members only possess domain 3. Antiapoptotic and proapoptotic family members may contain a C‐terminal ‘‘anchor’’ sequence that targets them to membranes


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some Bcl‐2 family proteins may not allow their insertion into the membrane until after the protein is activated during apoptosis (Chan and Yu, 2004). Bcl‐2 family members that lack this C‐terminal sequence are diffusely distributed in the cytoplasm. Almost all Bcl‐2 family members form homo‐ or heterodimers, and interactions between them are mediated by their Bcl‐2 homology domains (BH1–BH4) (Zha et al., 1996a; Sattler et al., 1997; Adams and Cory, 1998, 2001; Huang and Strasser, 2000; Lindenboim et al., 2001; Cory and Adams, 2002). The antiapoptotic members of the Bcl‐2 family share sequence homology in all four domains (> Figure 17-3), but the BH4 region appears to be important for their antiapoptotic functions (Hunter et al., 1996; Cheng et al., 1997; Huang et al., 1998). Although not without exception (see below), the BH4 region is primarily confined to this subclass of family members. The BH3‐only proapoptotic family members, such as Bid, Bik, Bad, Bim, Noxa, Puma, Hrk, and Spike, contain only the BH3 domain and share sequence homology in that region (> Figure 17-3). This domain appears to be necessary and sometimes sufficient for binding to antiapoptotic family members and promoting cell death (Chittenden et al., 1995; Zha et al., 1996a; Sattler et al., 1997; Kelekar and Thompson, 1998; Lindenboim et al., 2001; Chan and Yu, 2004). These proapoptotic family members act as upstream ‘‘sensors’’ for various apoptotic stimuli (Huang and Strasser, 2000; Cheng et al., 2001; Letai et al., 2002; Marani et al., 2002; Chan and Yu, 2004). For example (> Figure 17-4) growth factor deprivation causes dephosphorylation of Bad, which is then free to translocate to mitochondria where it can antagonize the antiapoptotic actions of Bcl‐xL (Zha et al., 1996b). Upon serum withdrawal or cytokine removal, Bim is released from microtubules and translocates to mitochondria to inhibit Bcl‐2 (Puthalakath et al., 1999). Following DNA damage, the transcription factor p53 induces transcription of Noxa and Puma (p53 upregulated modulator of apoptosis) which then translocate to mitochondria to inhibit the antiapoptotic Bcl‐2 family members (Oda et al., 2000; Nakano and Vousden, 2001; Yu et al., 2001; Yakovlev et al., 2004). Death receptor activation causes caspase‐8‐mediated cleavage of Bid, which then targets the mitochondria to enhance the function of other proapoptotic Bcl‐2 family members (Li et al., 1998; Luo et al., 1998; Zha et al., 2000). The Bax‐like proapoptotic Bcl‐2 family members, which includes Bax, Bok, and Bak, generally lack the BH4 domain and share sequence homology in the other three domains (> Figure 17-3). One exception is Bcl‐xS, which contains a BH4 domain, but lacks BH1 and BH2 (Lindenboim et al., 2001). In fact, Bcl‐xS is the only known proapoptotic family member that contains a strong BH4 domain. Members of this subfamily generally participate downstream from the BH3‐only family members in the apoptotic cascade and require an activation event before they can promote apoptosis (Cheng et al., 2001). For example, the voltage‐dependent ion channel 2 (VDAC2) associates with Bak at the outer‐mitochondrial membrane to keep it in an inactive conformation in healthy cells. In the presence of an apoptotic stimulus, truncated Bid (tBid), Bad, or Bim (BH3‐only family members) can displace VDAC2, allowing Bak to oligomerize and participate in apoptosis (Cheng et al., 2003). Under homeostatic conditions, antiapoptotic Bcl‐2 family members (e.g., Bcl‐2, Bcl‐xL) block apoptosis at the mitochondria by preventing the release of proapoptotic factors into the cytoplasm (Kluck et al., 1997; Yang et al., 1997), and the BH3‐only family members are held ‘‘in check’’ by a variety of mechanisms (e.g., phosphorylation, cytoskeletal sequestration). The presence of an apoptotic stimulus activates the appropriate BH3‐only proapoptotic family members, which remove the inhibition at the mitochondria. Actions of the BH3‐only family members enables the Bax‐like proapoptotic family members to move from the cytoplasm to the mitochondria (Wolter et al., 1997; Putcha et al., 1999), where they can initiate the release of the proapoptotic factors (Kluck et al., 1997; Jurgensmeier et al., 1998; Scorrano and Korsmeyer, 2003; Yamaguchi et al., 2003) (> Figure 17-4). In fact, Bax/Bak participation is required for subsequent activation of caspases in many forms of apoptotic death (Wei et al., 2001; Zong et al., 2001; Ruiz‐Vela et al., 2005). While some evidence suggests that cytochrome c release is passive and is the result of mitochondrial swelling and rupture of the outer‐mitochondrial membrane (Vander Heiden et al., 1997), other evidence suggests that Bcl‐2 family members actually permeabilize the outer‐mitochondrial membrane. Although the mechanisms of permeabilization are not clear, two prominent theories exist. First, Bcl‐2 proteins may actually form channels in the outer‐mitochondrial membrane. In support, Bax (Antonsson et al., 1997; Schlesinger et al., 1997; Korsmeyer et al., 2000), Bak (Korsmeyer et al., 2000), Bcl‐2 (Schendel et al., 1997;

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. Figure 17-4 This diagram illustrates some of the apoptotic stimuli that initiate apoptosis by activating upstream BH3‐only proapoptotic family members, which then propagate the death signal to other proapoptotic family members. A black ‘‘X’’ indicates a protein or event that is inhibited by the preceding component

Schlesinger et al., 1997), cleaved Bcl‐xL (Basanez et al., 2001), full‐length Bcl‐xL (Minn et al., 1997), and tBid (Schendel et al., 1999) can form channels in artificial membranes. Caspase‐8 was the first protease shown to cleave Bid (Li et al., 1998), which suggests that activation of upstream caspases may be required for some Bcl‐2 proteins to participate in pore formation. However, Bid cleavage can occur in a caspase‐independent manner by the serine protease granzyme B (Barry et al., 2000; Heibein et al., 2000; Alimonti et al., 2001), lysosomal proteases (Stoka et al., 2001; Reiners et al., 2002), or calpains (Chen et al., 2001; Mandic et al., 2002). Second, Bcl‐2 family proteins may permeabilize the mitochondrial membrane by interacting with preexisting channels, such as the permeability transition pore (Marzo et al., 1998; Zamzami et al., 2000; Antonsson, 2001; Zamzami and Kroemer, 2001; Tsujimoto and Shimizu, 2002).


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3.3 The Death Receptor (Extrinsic) Pathway of Apoptotic Cell Death The extrinsic pathway of apoptotic cell death is a death receptor‐mediated pathway, which is initiated by ligation of a transmembrane death receptor of the TNF/nerve growth factor superfamily. Members of this death receptor family include TNFR‐1 (p55, CD120), Fas (Apo1, CD95), death receptor‐3 (Apo3, WSL‐1, TRAMP, LARD), death receptor‐4 (TRAIL‐R1), and death receptor‐5 (Apo2, TRAIL‐R2, TRICK2, KILLER). For example, binding of Fas ligand (FasL) to the Fas receptor leads to clustering of the receptors’ intracellular DDs. Clustered receptors form receptor microaggregates at the cell surface, which produce a conformational change that enables the adapter protein, Fas‐associated protein with death domain (FADD; also known as the mediator of receptor‐induced toxicity (MORT1)), to bind to the clustered receptors through its DD (> Figure 17-5). Similarly, binding of TNF to the TNFR causes clustering of the . Figure 17-5 This diagram highlights some of the participants in the cell‐surface death receptor‐mediated (extrinsic) apoptotic pathway. A black ‘‘X’’ indicates a protein or event that is inhibited by the preceding component

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receptors’ intracellular DDs at the cell surface. TTRADD is then recruited to the receptor aggregates via its DD to serve as an intermediate adapter for the TNFR complex and FADD. It is important to note that recruitment of TRAF‐2 and RIP to the TNFR complex can lead to cell survival (proinflammatory pathway), whereas recruitment of FADD can lead to cell death. Cell fate following activation of the TNFR is dependent on the level of activation of these alternate pathways. During apoptosis, the complex formed between the receptor microaggregates and the adapter proteins at the cytoplasmic surface of the extracellular membrane is called the death‐inducing signaling complex (DISC). FADD recruits procaspase‐8 or ‐10 monomers to the DISC via homophilic interactions with their N‐terminal DEDs (> Figure 17-5) (Kischkel et al., 2001; Wang et al., 2001). Induced proximity enables dimer formation and the initiator caspases are activated through autoproteolysis, which causes removal of their N‐terminal DEDs and release into the cytoplasm (Muzio et al., 1998; Salvesen and Dixit, 1999; Boatright et al., 2003; Chang et al., 2003). The activated caspases can then cleave and activate downstream executioner caspases (Stennicke et al., 1998), which may include caspase‐12 (Wang et al., 2003). Activated caspase‐8 may also translocate to the nucleus of apoptotic cells to act as an executioner caspase to cleave poly(ADP‐ribose) polymerase‐2 (PARP‐2) (Benchoua et al., 2002). Activation of the extrinsic pathway alone does not kill some cell types and these cells also require the release of proapoptotic stimuli from mitochondria (Kluck et al., 1997; Scaffidi et al., 1998) to amplify the death signal. In these cases, activated caspase‐8 cleaves Bid (Li et al., 1998; Luo et al., 1998) and the C‐terminal fragment of tBid targets mitochondria to promote the release of apoptotic factors such as cytochrome c (Luo et al., 1998). Caspase‐8 activation is also responsible for cellular acidification prior to apoptosis, which is necessary for mitochondrial dysfunction, cytochrome c release, and caspase‐9 activation (Liu et al., 2000). Because caspase‐12 has been implicated in ER stress‐induced apoptosis (Nakagawa and Yuan, 2000; Nakagawa et al., 2000; Rao et al., 2001, 2002a, b), activation of death receptors may also trigger apoptotic mechanisms at the ER to amplify the death signal.

3.4 The ER (Intrinsic) Pathway of Apoptotic Cell Death A second ‘‘intrinsic’’ pathway of apoptotic cell death has recently been discovered and involves the ER. The ER is well‐known for its functions in lipid synthesis, protein folding and assembly, as well as storing intracellular calcium (Ca2þ) (Gething and Sambrook, 1990; Berridge, 2002; Michalak et al., 2002; Groenendyk et al., 2004). ER stress occurs under conditions that disrupt normal ER functions and cause malfolded proteins to accumulate in the ER lumen (e.g., disruption of intracellular Ca2þ homeostasis, disruption of protein glycosylation). Increases in unfolded or malfolded proteins trigger the unfolded protein response (UPR), which involves cell cycle arrest, attenuation of protein synthesis, induction of ER‐ localized chaperone proteins and folding catalysts, and activation of ER‐associated protein degradation (Kozutsumi et al., 1988; Brewer and Diehl, 2000; van Laar et al., 2001; Rao et al., 2004). If the ER stress cannot be resolved and cellular homeostasis is not reestablished, the cell dies by apoptosis (Welihinda et al., 1999; Kaufman, 2002; Rao et al., 2004). The mechanisms of ER stress‐induced apoptosis are not well understood, but there appears to be two pathways that may be initiated following ER stress (> Figure 17-6). One pathway is dependent on caspase‐ 12, which is localized to the cytosolic face of the ER (Nakagawa and Yuan, 2000; Nakagawa et al., 2000; Rao et al., 2001, 2002a, b). Several models have been proposed to explain caspase‐12 activation. The first model proposes that during ER stress misfolded proteins bind to the chaperone protein Grp78/Bip, which causes Grp78/Bip to release the ER stressor protein Ire1a. Ire1a then oligomerizes and autoactivates (Bertolotti et al., 2000). Although a trimeric complex has not been confirmed, activated Ire1 can recruit TRAF2 (Urano et al., 2000), which could release procaspase‐12 from TRAF‐2 and cause procaspase‐12 to homodimerize and autoactivate (Yoneda et al., 2001). Grp78/Bip may also bind to procaspases‐7 and ‐12 at the ER surface under homeostatic conditions and then release them into the cytoplasm during apoptosis (Rao et al., 2002b; Reddy et al., 2003). The second model proposes that Ca2þ released from the ER into the cytoplasm during ER stress may activate cytoplasmic calpain, which could translocate to the ER and cleave


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. Figure 17-6 This diagram highlights some of the participants in the endoplasmic reticulum (ER) stress‐induced (intrinsic) apoptotic pathway. A black ‘‘X’’ indicates a protein or event that is inhibited by the preceding component

the prodomain of procaspase‐12 to activate the caspase (Nakagawa and Yuan, 2000). The third model proposes that caspase‐7 may translocate from the cytoplasm (Chandler et al., 1998) to the ER, where it could cleave procaspase‐12 to generate the activated caspase (Rao et al., 2001). Because caspase‐12 is an initiator and caspase‐7 is an executioner, this event may serve to amplify the death signal of activated caspase‐12. Although the precise mechanisms are currently unknown, Zong et al. (2003) found evidence to suggest that caspase‐12 activation was dependent on ER‐targeted Bak. Once activated, caspase‐12 may process downstream caspases in the cytoplasm or target other unidentified substrates. Some data suggests that activated caspase‐12 can cleave and activate caspase‐9 in a cytochrome c‐ and Apaf‐1‐independent manner, which could cause subsequent activation of executioner caspases (Rao et al., 2001, 2002a; Morishima et al., 2002). Although Hitomi et al. (2003) also found evidence that caspase‐12 indirectly activated caspase‐3 during ER stress, they did not find evidence of caspase‐9 activation.

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Gene expression experiments recently suggested that human caspase‐12 has a genetic mutation that renders it nonfunctional (Fischer et al., 2002; Lamkanfi et al., 2004). However, several human cell lines express caspase‐12 protein (Nakagawa et al., 2000; Bitko and Barik, 2001; Rao et al., 2001; Yoneda et al., 2001; Xie et al., 2002). Although procaspase‐12 is processed during ER stress‐induced apoptosis, activated caspase‐12 may be dispensable for the execution of cell death (Lamkanfi et al., 2004). Additional work needs to be conducted to determine if caspase‐12 plays a major role in ER stress‐induced apoptosis in humans. The second pathway activated by ER stress is caspase‐12‐independent/mitochondria‐dependent and is strongly regulated by Bcl‐2 family members (Hacki et al., 2000; McCullough et al., 2001; Nutt et al., 2002b; Scorrano et al., 2003; Thomenius et al., 2003). Alterations in Ca2þ homeostasis are known to induce apoptosis (Nakamura et al., 2000; Ferrari et al., 2002; Nutt et al., 2002a; Scorrano et al., 2003; Smaili et al., 2003) and may reinforce cross‐talk between the ER and mitochondria (Hacki et al., 2000; Boya et al., 2002; Hori et al., 2002; Mattson and Chan, 2003), which are located in close proximity to one another (Rizzuto et al., 1998). Bap31 is an ER transmembrane protein that binds nascent proteins in transit between the ER and Golgi complex. Under homeostatic conditions, Bap31 forms a complex with antiapoptotic Bcl‐2 family members (Bcl‐2 or Bcl‐xL) and the long isoform of procaspase‐8 (procaspase‐8L) at the cytoplasmic face of the ER (> Figure 17-6) (Ng et al., 1997; Breckenridge et al., 2002). BH3‐only family members, such as Spike, may promote apoptosis at the ER by binding to Bap31 and preventing its association with antiapoptotic family members (Mund et al., 2003). Bap31 is a preferred substrate for initiator caspases‐1 and ‐8 (Ng et al., 1997) and is cleaved into a membrane‐integrated fragment (p20) during apoptosis. Bap31 cleavage as well as Bax/Bak oligomerization during ER stress are important events for release of ER Ca2þ and propagation of the apoptotic stimulus to mitochondria (Nguyen et al., 2000; Nutt et al., 2002a, b; Breckenridge et al., 2003; Scorrano et al., 2003). Although the precise mechanisms are not known, transcription of Puma also occurs during ER stress and is correlated with subsequent cytochrome c release from mitochondria and caspase activation (Reimertz et al., 2003). Release of Ca2þ from the ER can cause cell death by directly affecting mitochondrial function, as well as activating the Ca2þ‐dependent protease calpain (Wang, 2000). As already mentioned, activated calpain in the cytoplasm can activate caspase‐12 at the ER (Nakagawa and Yuan, 2000). Calpain can also cleave the calcineurin inhibitor cain/cabin 1 resulting in the activation of calcineurin (Kim et al., 2002). Activated calcineurin can lead to Bad dephosphorylation and subsequent caspase‐3 activation (Wang et al., 1999; Springer et al., 2000). Calpain can also cleave Bax and Bid to activate these proapoptotic proteins (Wood et al., 1998; Choi et al., 2001; Mandic et al., 2002). Thus, calpains and caspases may cooperate to execute apoptosis following ER stress.

3.5 Additional Pathways of Caspase Activation Although the mechanisms are not completely understood, caspase‐2 is activated in response to cellular stress and occurs in the early stages of apoptosis, upstream of mitochondrial events (Mancini et al., 2000; Troy et al., 2001; Lassus et al., 2002; Robertson et al., 2002; Tinel and Tschopp, 2004). Caspase‐2 has been localized to mitochondria, Golgi complex, cytoplasm, and nucleus (Zhivotovsky et al., 1999; Mancini et al., 2000; Paroni et al., 2002). Procaspase‐2 undergoes autocatalytic cleavage following dimerization for its initial activation (Read et al., 2002; Baliga et al., 2004; Ho et al., 2005). Processing by caspase‐3 (Li et al., 1997a; O’Reilly et al., 2002) amplifies its cell death activity (Baliga et al., 2004). Unlike most initiator caspases, activated caspase‐2 appears inactive toward most caspase zymogens (Guo et al., 2002), except possibly procaspase‐7 (Ho et al., 2005), and it is not inhibited by IAPs (Ho et al., 2005). Procaspase‐2 is the only caspase localized to the nucleus and nuclear localization is dependent on its prodomain (Colussi et al., 1998). Nuclear caspase‐2 is activated following DNA damage and causes release of mitochondrial proapoptotic factors and subsequent activation of caspases‐9 and ‐3 (Guo et al., 2002; Lassus et al., 2002; Paroni et al., 2002; Robertson et al., 2002, 2004). Although relocalization from the nucleus to the cytoplasm is not necessary for activated caspase‐2 to promote mitochondrial release of proapoptotic factors (Paroni et al., 2002; Robertson et al., 2002), recent evidence suggests that activated caspase‐2 can actually permeabilize mitochondrial membranes and disrupt the association of cytochrome c with cardiolipin (Enoksson et al.,


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2004), which normally binds cytochrome c to the inner‐mitochondrial membrane and must be disrupted for cytochrome c release (Ott et al., 2002). Together, these findings suggest an important link between nuclear stress and the mitochondrial pathway of apoptosis. In addition to its activation in response to DNA damage, caspase‐2 may also be activated during death receptor‐mediated apoptosis (see Lassus et al., 2002). The caspase and RIP adapter with DD (CRADD; also known as RAIDD) has an N‐terminal CARD that can interact with procaspase‐2 and a C‐terminal DD that can interact with RIP, which enables CRADD to serve as an adapter protein to recruit procaspase‐2 to the TNFR complex (Ahmad et al., 1997; Duan and Dixit, 1997; Ashkenazi and Dixit, 1998; Chou et al., 1998). Ceramide‐induced apoptosis revealed that caspase‐2 participates upstream of caspase‐8 activation and mitochondrial damage (Lin et al., 2004). In some cells, caspase‐2 activation is required for efficient cleavage of Bid following activation of death receptors‐4 and ‐5 (Wagner et al., 2004). Caspase‐2 can also be recruited into a complex with TRAF2 and RIP via its CARD, activating proinflammatory pathways (Lamkanfi et al., 2005). These findings suggest that caspase‐2 may participate in cell death or cell survival following activation of death receptors.

4

Substrate Cleavage by Activated Caspases

The intrinsic and extrinsic apoptotic pathways converge on a final common pathway where executioner caspases are activated to cleave other caspases and amplify the death signal, as well as to cleave numerous cellular substrates to dismantle the cell. Cleavage of specific cellular substrates has been linked to DNA fragmentation and particular features of the apoptotic morphology (Thornberry and Lazebnik, 1998; Saraste and Pulkki, 2000). Activated caspases can (1) inactivate proteins that prevent apoptosis, (2) alter cell structure by cleaving cytoskeletal and nuclear proteins, and (3) inactivate proteins involved in repair. For example, the antiapoptotic proteins, Bcl‐2 and Bcl‐xL, are cleaved by caspase‐3 to generate C‐terminal fragments that are proapoptotic (Cheng et al., 1997; Clem et al., 1998; Fujita et al., 1998; Kirsch et al., 1999). Caspases may also cleave proapoptotic Bcl‐2 family proteins to make them more potent inducers of apoptosis (Condorelli et al., 2001). Caspase cleavage of structural and nuclear proteins, such as gelsolin (Kothakota et al., 1997), p21‐activated kinase 2 (PAK2; Rudel and Bokoch, 1997), and lamin A (Orth et al., 1996; Takahashi et al., 1996a, b) cause membrane blebbing and apoptotic bodies (see > Figure 17-2). As already mentioned, executioner caspases cleave ICAD (also known as DNA fragmentation factor‐45 (DFF45)), which frees CAD (DFF40) to translocate to the nucleus and digest DNA (Liu et al., 1997a; Enari et al., 1998; Sakahira et al., 1998). Caspases also cleave and activate acinus, which causes chromatin condensation (Sahara et al., 1999). PARP‐1 is a nuclear and mitochondrial enzyme activated by DNA damage that has varied functions in DNA repair, transcription and replication, as well as cell differentiation, proliferation, and death (Burzio et al., 1981; Masmoudi et al., 1988; Lautier et al., 1993; D’Amours et al., 1999; Shall and de Murcia, 2000; Virag and Szabo, 2002; Du et al., 2003). During apoptosis, PARP is cleaved by caspases (Kaufmann et al., 1993; Lazebnik et al., 1994; Casciola‐Rosen et al., 1996; Takahashi et al., 1996a), which helps preserve energy stores but prevents DNA repair. Caspases can also cleave the catalytic subunit of the DNA‐dependent protein kinase (DNA‐PKcs) (Casciola‐Rosen et al., 1996), which is important in repairing double‐stranded DNA breaks. Although these are only a few examples, almost 300 caspase substrates have now been identified (Fischer et al., 2003). Caspases generally recognize a tetrapeptide sequence and cleave their substrates after particular aspartate residues. Specifically, an aspartate must reside in the first position of the recognition sequence (Thornberry et al., 1997; Garcia‐Calvo et al., 1998). However, the sequence of residues that follows the initial aspartate determines the specificity of substrate cleavage by particular caspases. After analyzing caspases‐1 to ‐9, Thornberry et al. (1997) determined that the amino acid in the fourth position conferred the greatest specificity. The mammalian caspases‐1 through ‐10 are the best studied and can be subdivided into three groups according to their preferred amino acid recognition sequence (Thornberry et al., 1997; Garcia‐Calvo et al., 1998). Caspases‐1, ‐4, and ‐5 cleave their substrates after the amino acid sequence (W/L) ExD. Caspases‐2, ‐3, and ‐7 cleave after the sequence DExD. Caspases‐6, ‐8, ‐9, and ‐10 prefer the sequence (I/L/V)ExD.

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Caspase‐Independent Apoptosis

Caspase‐independent pathways refer to those cascades of events that cause DNA fragmentation and some of the morphological features of apoptosis in the absence of significant caspase activation. The actions of AIF and Endo G are thought to exemplify caspase‐independent apoptotic processes in mammalian cells through their actions at the nucleus following release from the mitochondria (Susin et al., 1999b; Daugas et al., 2000b; Li et al., 2001). Although less fully characterized, HtrA2/Omi (Suzuki et al., 2001) and other mitochondrial and cytoplasmic proteins (Lorenzo and Susin, 2004) may have caspase‐independent functions in cell death. Activation of caspase‐independent pathways does not mean that caspases are not eventually activated. In several models, caspase‐independent apoptosis causes cells to die with only partial condensation of nuclear chromatin, with or without DNA fragmentation, while the other features of apoptosis are generally maintained (Miller et al., 1997; Susin et al., 1999b, 2000; Doonan et al., 2003). Subsequent activation of caspase‐dependent apoptotic pathways may be required for the complete and efficient transformation of the dying cell. In support, Susin et al. (1999b) found that AIF can cause caspase‐9 activation. However, the notion that AIF and Endo G are mediators of caspase‐independent apoptosis contradicts findings suggesting that mitochondrial release of AIF and Endo G is caspase‐ dependent (Arnoult et al., 2002, 2003; Cregan et al., 2002). The apparent contradiction in the release of these mitochondrial proapoptotic factors needs further investigation, but probably depends on the type of cell and the particular apoptotic stimulus (see Cregan et al., 2004 for discussion). Because the focus of this chapter is on the role of caspases in cell death, these alternative death pathways will not be discussed further.

6

Non‐Apoptotic Functions of Caspases

Evidence suggests that caspase activation does not always equal cell death (Zeuner et al., 1999). Such findings are not reviewed in detail as it is beyond the scope of this chapter. Caspase‐like proteases have a role in the proliferation and differentiation of several cell types (Ishizaki et al., 1998; Fernando et al., 2002; Sordet et al., 2002; Schwerk and Schulze‐Osthoff, 2003) and in the activation of T cells (Miossec et al., 1997; Wilhelm et al., 1998; Wu et al., 1999; Salmena et al., 2003). Because caspases process several proteins involved in cell cycle control, caspases may regulate both life and death of a cell (Zeuner et al., 1999). Therefore, it is important to determine whether caspases activated following injury or disease are affecting cell survival or cell death.

7

The Role of Caspases in Neurotrauma

7.1 Introduction Since all of our cells carry the machinery necessary to conduct its own demise, it is important to determine whether dysregulation of the cell death machinery underlies the cell death that occurs after nervous system trauma. Early investigations suggested that caspase inhibition offered neuroprotection in some experimental models (Milligan et al., 1995; Deshmukh et al., 1996; Armstrong et al., 1997; Eldadah et al., 1997; Cutillas et al., 1999; Li et al., 2000; Cao et al., 2002; Knoblach et al., 2002), and deletion of single caspases reduced cell death in development (Kuida et al., 1996). However, other data suggests that caspase inhibition only delays apoptosis (McCarthy et al., 1997; Miller et al., 1997; Stefanis et al., 1999; Keramaris et al., 2000; Selznick et al., 2000; Cregan et al., 2002). Others have reported that caspase inhibition does not prevent cell death, but alters the form of the demise (Xiang et al., 1996; Hirsch et al., 1997; Sarin et al., 1997), which may depend on cellular ATP levels (Eguchi et al., 1997; Leist et al., 1997, 1999; Stridh et al., 1999). Still others have reported that caspase inhibition can exacerbate cell death (Vercammen et al., 1998; Cauwels et al., 2003). These apparent discrepancies may partially be due to the criteria used to assess cell death or cytoprotection. When mammalian cells are treated with caspase inhibitors such as N‐benzyloxy‐carbonyl‐ Val‐Ala‐Asp‐fluoromethylketone (zVAD‐fmk), suppression of caspases appears to inhibit apoptosis when chromatin condensation and DNA fragmentation are the criteria for defining cell death. However, when


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other signs of cell death such as mitochondrial dysfunction, phosphatidylserine exposure, or plasma membrane permeabilization are used to define cell death, then caspase inhibition does not appear cytoprotective (Perfettini and Kroemer, 2003). Therefore, it is important to understand the circumstances under which caspases participate in cell death, the form of the death, and whether caspase inhibition will salvage cells or simply divert or delay cell death.

7.2 Ischemic Brain Injury (Stroke) The involvement of apoptotic cell death in CNS injury has been studied extensively with models of focal or global cerebral ischemia (Kirino, 1982; Pulsinelli et al., 1982; Choi, 1996), and evidence suggests that several members of the caspase family are activated in these models (Loddick et al., 1996; Friedlander et al., 1997; Hara et al., 1997; Chen et al., 1998; Namura et al., 1998; Schielke et al., 1998). For example, Asahi et al. (1997) reported that caspase‐2 mRNA increased at 4–16 hr and caspase‐3 mRNA increased at 16–24 hr after permanent middle cerebral artery occlusion (MCAO) in adult rats. In contrast to rats, Bergeron et al. (1998) found no change in caspase‐2 protein levels after transient or permanent MCAO in mice. In a more extensive study, Harrison et al. (2001) found a large increase in caspase‐11 mRNA in the ipsilateral cortex 3–6 hr following permanent MCAO in adult rats which continued for 24 hr, whereas caspase‐6 mRNA peaked at 6 hr postinjury. Although mRNA for caspases‐3, ‐7, and ‐8 was elevated by 6 hr postinjury, the largest increase was between 12 and 24 hr, which was followed by increased caspase‐1 mRNA at 24 hr postischemia. In contrast to the other caspases, caspase‐2 mRNA did not change significantly in the ipsilateral cortex during the first 24 hr, and caspase‐9 mRNA decreased slightly by 12 hr postinjury. Following temporary MCAO in adult rats, activated caspase‐3 appeared in neuronal perikarya and dendrites in the territory of the middle cerebral artery within minutes of reperfusion and was followed by caspase‐3‐like enzyme activity and DNA fragmentation (Namura et al., 1998). Recent evidence suggests that activated caspase‐12 may also have a role in cell death following MCAO (Mouw et al., 2003; Shibata et al., 2003). Time‐dependent changes in caspase activation have also been reported with models of global ischemia. Chen et al. (1998) found that activated caspase‐3 was upregulated in the hippocampus and striatum from 8 to 72 hr, and activated caspase‐1 modestly increased in hippocampal lysates of adult rats at 72 hr postinjury. Caspase‐3 mRNA was upregulated in the CA1 region of the hippocampus 24 hr after four‐ vessel occlusion, and then increased up to 72 hr postinjury. No change in caspase‐2 expression was observed (Ni et al., 1998). Following transient bilateral carotid artery occlusion in rats, Ouyang et al. (1999) reported that cytochrome c release into the cytoplasm was apparent by 36 hr and DEVD‐cleaving activity increased to twice the control levels by 48 hr postischemia. Although different features of the apoptotic cascade were examined in each study, these findings collectively illustrate a time‐dependent evolution of caspase activity and cell death following ischemic injury in vivo. Discrepancies in the types of caspases activated and their time course are probably due to the specifics of the injury model utilized (species, strain, type, and duration of injury, etc). Therefore, further research is needed to fully characterize the spatiotemporal evolution of caspase activity and cell death following ischemic injury. The importance of such work is illustrated by the fact that activated caspase‐3 was observed in brain neurons of human patients who died 12 hr to 9 days following cardiac arrest with resuscitation (Love et al., 2000).

7.3 Traumatic Brain Injury Evidence from several laboratories supports the hypothesis that apoptotic cell death in the rat brain occurs following traumatic injury (i.e., lateral fluid percussion or cortical contusion injuries; see Raghupathi et al., 2000). Alan Faden’s laboratory presented some of the first evidence that apoptosis in traumatic brain injury (TBI) was dependent, in part, upon caspase‐3 activation (Yakovlev et al., 1997). In this study, DNA fragmentation was observed in cytosolic extracts isolated from the ipsilateral cortex and hippocampal

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formation as early as 4 hr following lateral fluid percussion injury in rats. Caspase‐3 mRNA increased four‐ to fivefold at this time‐point and was accompanied by a 25–50% increase in caspase‐3‐like enzyme activity. Caspase‐1 mRNA also increased after trauma, but to a lesser extent than caspase‐3 mRNA. In a similar model, Knoblach et al. (2002) found increased activated caspase‐3 and ‐9, which often persisted for 2 days postinjury. These authors reported little change in activated caspase‐8. Using a controlled cortical impact model of TBI, Beer et al. (2001) reported an increase in activated caspase‐8 and ‐3 in the ipsilateral cortex 6–72 hr postcontusion, with peaks at 24 hr and 48 hr, respectively. Recent evidence from a rat model of controlled cortical impact has also implicated caspase‐12 in the pathology of brain trauma (Larner et al., 2004). A biochemical hallmark of TBI (and other traumatic CNS events) is the breakdown of certain cytoskeletal proteins critical for maintaining structural integrity. One such protein is a‐spectrin, which has specific amino acid sequences that allow it to be processed by at least two different cysteine proteases: calpain or activated caspase‐3. Following TBI, Hayes and colleagues (Posmantur et al., 1998; Pike et al., 1998a, b) reported that a‐spectrin was primarily processed by calpain in the injured cortex, while the caspase‐3‐processed form of a‐spectrin predominated deeper structures (hippocampus and striatum). This observation is important because tissue loss in the hippocampus and striatum is less severe than the tissue loss suffered by the overlying cortex. McIntosh and coworkers reported that ICAD/DFF45 is cleaved in a caspase‐3‐mediated manner within 2 hr following TBI (Zhang et al., 1999). As a consequence, CAD/DFF40 would then be released from its heterodimer inhibitory complex and enter the nucleus to degrade chromosomal DNA (Enari et al., 1998). Taken together, the results of these studies provide additional support that caspase‐3 activation and substrate degradation are major factors responsible for apoptotic cell death following experimental TBI. The importance of these observations is exemplified by reports of activated caspase‐8 (Zhang et al., 2003), caspase‐1, caspase‐3, and DNA fragmentation (Clark et al., 1999) in the human brain following traumatic injury. Caspase‐3‐like activity has also been observed in the cerebrospinal fluid of TBI patients (Harter et al., 2001).

7.4 Traumatic Spinal Cord Injury A number of studies have provided compelling evidence of widespread apoptosis of neurons, oligodendroglia, and microglia following spinal cord injury (SCI) (Katoh et al., 1996; Li et al., 1996; Crowe et al., 1997; Liu et al., 1997b; Shuman et al., 1997; Emery et al., 1998; Hayashi et al., 1998; Lou et al., 1998; Yong et al., 1998). An excellent review by Beattie and colleagues provides a summary of the current findings (Beattie et al., 2000). Many of the cells with an apoptotic phenotype were located several millimeters from the injury epicenter, indicating that this cell death process contributes to the pattern of white and gray matter tissue loss commonly found in traumatic SCI. At least one group has hypothesized that apoptosis of oligodendroglia in areas distant to the injury epicenter contributes to long‐term neurological dysfunction (Crowe et al., 1997; Shuman et al., 1997). Specifically, the loss of oligodendroglia can result in demyelination and dysfunction of uninjured descending axonal tracts that control voluntary motor function (Blight, 1983, 1989; Blight and Decrescito, 1986). Therefore, understanding the intracellular signaling pathway(s) involved in this cell death process may prove critical for developing therapies that will reduce oligodendroglial cell loss following SCI. In 1998, cells with apoptotic morphologies were observed in injured human spinal cord tissue, and these cells contained the activated form of caspase‐3 (Emery et al., 1998). Around the same time, our laboratory was investigating the components of the caspase‐3 apoptotic cascade using an experimental model of traumatic SCI in rats (Springer et al., 1999). Within 1 hr following injury, caspase‐3 enzyme activity levels increased sixfold and remained elevated up to 24 hr in cytosolic extracts from the injured spinal cords. The results of these initial experiments led us to speculate that the molecular pathways responsible for caspase‐3 activation in vitro (i.e., cytochrome c release and caspase‐9 activation) should be detectable in the injured spinal cord. Using immunoblotting and semiquantitative densitometry, we found that cytochrome c levels were significantly elevated by 30 min after injury, and remained elevated for at least 24 hr. As predicted, caspase‐9 activation followed a temporal pattern similar to the pattern observed for cytochrome c release. Similar to findings


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with TBI (Zhang et al., 1999), there was clear evidence of DFF45/ICAD cleavage at 1 and 4 hr following SCI (Springer et al., 1999). While the pattern of DFF45/ICAD proteolysis is consistent with a role for activated caspase‐3, other possibilities including the contribution of other caspases (e.g., caspase‐7) that also cleave DFF45/ICAD as well as caspase‐independent apoptotic cell death cannot be ruled out. Nevertheless, these results provide compelling evidence that apoptotic cell death following SCI involves the proteolytic actions of activated caspase‐3, and occurs downstream of activated caspase‐9. These findings have recently been supported and extended by other investigators. For example, Festoff and colleagues documented increased caspase‐3 expression and activity in the injured rat spinal cord (Citron et al., 2000). These investigators reported a rapid upregulation of caspase‐3 mRNA and protein in regions (mRNA) and cells (protein) of the spinal cord with the highest amounts of apoptosis postinjury. With a mouse model of SCI, Takagi et al. (2003) examined caspase activation between 8 hr and 7 days postinjury and found that caspase‐8‐like activity was upregulated on days 3 and 7 and caspase‐3‐like activity was detected on day 7, when morphological changes to the cells were also apparent. In a more extensive study, Knoblach et al. (2005) examined the activation of caspases‐1, ‐2, ‐3, ‐6, ‐8, and ‐9 from 1 hr to 7 days following spinal cord contusion in rats. These authors reported that cleaved caspases‐3, ‐8, and ‐9 were detected in tissue homogenates 1–72 hr postinjury, and could be found several segments away from the point of impact as early as 4 hr postinjury. Furthermore, caspase‐3‐like activity was apparent as late as 7 days postinjury. Although these authors did not find evidence of caspase‐1, ‐2, and ‐6 activity, Li et al. (2000) previously detected activated caspase‐1 in a mouse model of SCI. The experimental findings presented above suggest that the rodent spinal cord contains the molecular machinery necessary for activation and execution of the caspase‐3 apoptotic pathway.

8

Therapeutic Interventions in CNS Trauma

8.1 Competitive Inhibitors The rapid onset of caspase activation in some injury models indicates that there is a relatively narrow therapeutic time window for implementing treatments that prevent caspase activation. However, apoptosis persists for hours to weeks following certain types of CNS injuries, so strategies that minimize or halt ongoing caspase activation and enzyme activity may minimize the cleavage of cellular substrates responsible for generating the apoptotic phenotype. Treatment with caspase inhibitors could reduce caspase activation and increase cell survival, as well as improve neurological outcome. In line with this, neuroprotective effects with administration of tetrapeptide‐based competitive substrates have been reported for the various models of CNS injury. Intracerebroventricular (ICV) administration of zVAD, a pan‐caspase inhibitor, reduced infarct volume in the brain of adult rats and mice by approximately 50% following a 2 hr MCAO, regardless of whether the inhibitor was administered 15 min prior to, or at the time of, reperfusion (Loddick et al., 1996; Hara et al., 1997). The decrease in infarct volume was observed at 18 hr postreperfusion and was sustained for 3 days. Furthermore, the treatment improved forepaw extension, circling, and walking at 18 hr following reperfusion (Hara et al., 1997). When the same authors administered the caspase‐3‐like tetrapeptide inhibitor, zDEVD‐fmk, to adult rats, infarct volume and behavioral deficits were reduced but to a lesser extent than when the pan‐caspase inhibitor was administered. ICV administration of zDEVD‐fmk reduced infarct volume by 30% following MCAO in adult mice (Hara et al., 1997), but did not reduce brain swelling (which may be due to caspase‐1 activation). When the middle cerebral artery was occluded for only 30 min, zVAD‐ fmk administration that was delayed for 6 hr after reperfusion still reduced infarct volume by 50%, which was as effective as drug pretreatment (Endres et al., 1998). In a mouse model of transient MCAO, Fink et al. (1998) demonstrated that zDEVD‐fmk was neuroprotective as long as it was administered prior to the appearance of caspase activity. Specifically, zDEVD‐fmk was neuroprotective when administered up to 9 hr after reperfusion, while the cleavage of DEVD‐containing substrates was low. Recent evidence suggests that the caspase‐1 (Rabuffetti et al., 2000; but see Chen et al., 1998) and caspase‐9 (Mouw et al., 2002) inhibitors may also be effective in promoting tissue preservation and functional recovery following ischemic injury.

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ICV administration of caspase inhibitors has also been effective with models of TBI and SCI. When zDEVD‐fmk was administered into the lateral ventricles at 30 min before and 6 and 24 hr after a lateral fluid percussion injury, caspase‐3 activation, DNA fragmentation, and the number of apoptotic neurons in the injured cortex were dramatically reduced. More importantly, the same treatment paradigm improved neurological function on several motor tasks sensitive to TBI (Yakovlev et al., 1997). These findings were supported by a separate study demonstrating that postinjury treatment with zDEVD‐fmk reduced both cell loss and lesion volume, but there was no evidence of improved neurological function (Clark et al., 2000). In contrast, Knoblach et al. (2002) administered a pan‐caspase inhibitor 15 min after lateral fluid percussion injury and found improved motor function and spatial learning. Li et al. (2000) administered zVAD‐fmk to mice immediately following spinal cord contusion and found reduced numbers of TUNEL‐positive cells and the amount of activated caspase‐1 and caspase‐3, and improved pain withdrawal and motor score in an open field. In a similar model, Takagi et al. (2003) administered a pan‐caspase inhibitor or a specific caspase‐3 inhibitor immediately after contusion to reduce caspase‐8‐ and caspase‐3‐like enzyme activities at 7 days postinjury. More recently, Colak et al. (2005) treated rats with the caspase‐9 inhibitor, zLEHD‐fmk, immediately after SCI and found a subsequent reduction in apoptosis at 24 hr and 7 days postinjury. Furthermore, rats showed improved functional outcome, as assessed on the inclined plane and a modified Tarlov motor grading scale. Knoblach et al. (2005) reported that intrathecal administration of the pan‐ caspase inhibitor, Boc‐d‐fmk, at 15 min postcontusion improved locomotor function on postinjury days 21 and 28. In contrast, treatment with the caspase‐3 inhibitor, zDEVD‐fmk, improved locomotor function on postinjury day 21 only. Together, these findings suggested that multiple caspases are involved in cell death and neurological dysfunction following CNS trauma. Caspase inhibitors that act upstream of activated caspase‐3 or that broadly inhibit multiple caspases may be most effective in tissue preservation and behavioral recovery following an insult. Although the results of these functional studies show promise, it will be important to determine whether these benefits are transient or long term.

8.2 Anti‐Apoptotic Genes Given our knowledge of the biochemical pathways involved in caspase activation, another putative therapeutic approach is to use replication‐defective viral vectors to deliver antiapoptotic genes to the injured CNS. This strategy was initially described as a way to limit cell death following ischemic injury (Martinou et al., 1994; Linnik et al., 1995). Certain viral vectors exhibit short‐term expression of the gene of interest (typically 3–6 weeks) due to nonintegration into the host genome (Smith and Romero, 1999; Kay et al., 2001). Transient expression of a transgene would be optimal in CNS injury because apoptosis occurs from hours to weeks following the initial insult, which has been well documented for SCI (Crowe et al., 1997; Liu et al., 1997b; Shuman et al., 1997). Expression of an antiapoptotic gene would not be required at extended time‐points when apoptosis is no longer a predominant event in the injured CNS. The genes of interest to be delivered to the injured CNS are based on their potential to inhibit different steps in the caspase‐3‐mediated apoptotic cascade. One approach would be to target events upstream to mitochondrial release of cytochrome c and Smac/Diablo. Therefore, overexpression of antiapoptotic members of the Bcl‐2 gene family, which bind to and inhibit the actions of proapoptotic Bcl‐2 family members, may be beneficial. Yukawa et al. (2002) delivered recombinant adenovirus carrying the human Bcl‐2 oncogene into the spinal cord of rats and found that adv‐Bcl‐2 injected immediately after injury suppressed lesion development at 72 hr postinjury, as assessed by a reduction in DNA fragmentation (TUNEL staining). When Zhao et al. (2003) delivered viral vectors that overexpressed Bcl‐2 to the infarct margin before focal ischemia in rats, neuron survival was improved and the cytosolic levels of cytochrome c and activated caspase‐3 were reduced. A second potential step that can be targeted in the caspase‐3 apoptotic cascade involves events downstream of cytochrome c release but upstream of caspase‐3 activation, such as the family of IAPs (Deveraux et al., 1998; Duckett et al., 1998; Ekert et al., 1999, 2001). Mammalian IAPs include XIAP, c‐IAP1, c‐IAP2, NAIP, and Survivin, but XIAP inhibits caspase‐3 activation at a Ki that is much lower than any of


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the other IAPs. Removal of XIAP inhibition of caspases is at least partly dependent upon Smac/Diablo (Srinivasula et al., 2000, 2001; Verhagen et al., 2000; Wu et al., 2000), which is a death factor released by mitochondria during apoptosis (Du et al., 2000; Verhagen et al., 2000). As previously discussed, Smac/ Diablo displaces XIAP from procaspase‐9 and procaspase‐3, allowing the caspases to become activated. Taken together, these studies suggest that overexpression of IAPs should reduce caspase‐3 activation in cells following CNS injury. In support, Xu et al. (1999) used adenoviral vectors to overexpress XIAP in the hippocampus of rats after four‐vessel occlusion and found improvements in neuron survival and spatial learning. Similarly, overexpression of NAIP reduced hippocampal damage postischemia (Xu et al., 1997).

8.3 Targeting the Mitochondria Strategies that target specific mitochondrial events may also prove beneficial as therapeutic interventions following CNS injury. Several experimental studies suggest that mitochondrial homeostasis is severely impaired following CNS injury and recent reports from our laboratory and others have demonstrated that mitochondrial permeability transition (mPT) occurs in models of TBI and SCI (Jin et al., 2004; Sullivan et al., 2005). Mitochondria play a critical role in the regulation of the caspase‐3 apoptotic cascade, and the pathophysiological role of mPT in apoptotic cell death indicates that inhibition of mPT should be effective toward reducing apoptotic cell death in vivo. One of the most potent inhibitors of mPT is cyclosporin A (CsA), an immunosuppressant commonly used for reducing organ transplant rejection. Once the connection was made between a potential role between mPT and apoptotic events, the use of CsA became a logical choice in attempting to reduce this form of cell death induced following CNS injury. In 1999, work from Povlishock’s group demonstrated that treatment with CsA reduced diffuse axonal injury and Ca2þ‐related cytoskeletal damage following TBI in rats (Okonkwo et al., 1999; Okonkwo and Povlishock, 1999). Scheff and Sullivan (1999) also demonstrated that CsA significantly reduce cortical damage (50%) in TBI when administered at 15 min and 24 hr postinjury. The immunosuppressive properties of CsA were most likely not responsible for its neuroprotective effects since neuroprotection was not observed when injured animals were treated with FK506, a potent immunosuppressant. Further, this same group (Sullivan et al., 1999) used several biochemical assays of mitochondrial integrity and bioenergetics to demonstrate that CsA treatment significantly attenuated mitochondrial dysfunction following TBI. CsA is FDA approved and is currently being used in clinical trials for TBI. However, CsA has significant dose‐dependent neurological effects, including seizures and cell death (Berden et al., 1985; de Groen et al., 1987; Walker and Brochstein, 1988; Famiglio et al., 1989; Kahan, 1989). Given the toxicity of high doses of CsA, the use of a nonimmunosuppressive CsA derivative that inhibits mPT and exhibits minimal toxicity would be attractive. We are currently testing a CsA derivative, NIM811, which has minimal toxicity compared to CsA (Waldmeier et al., 2002) in models of TBI and SCI. The use of NIM811 also allows one to directly test the hypothesis that mPT is critical to cell death induced by TBI or SCI and that the underlying mechanism of CsA‐mediated neuroprotection is not related to immunosuppression. In collaboration with Pat Sullivan, our preliminary results indicate that NIM811 increases tissue sparing, improves mitochondrial function, and reduces mitochondrial oxidative stress following TBI and SCI in adult rats. While these observations do not exclusively implicate mPT in cell death, these results are very supportive for a role of mPT modulation in the neuroprotective effects of CsA and its derivatives following CNS injury. Given these observations, the identification and development of pharmacological agents that target the mPT may prove beneficial in the acute treatment of traumatic CNS injury.

9

Conclusion

Most therapies discussed in the preceding sections were administered prior to, or at the time of, insult to maximize their neuroprotective potential. As such, these findings provide strong ‘‘proof of principle’’ that the caspase apoptotic cascades are at least partly responsible for the pathophysiological responses of the mammalian CNS to ischemic or traumatic insults. Because most individuals with traumatic CNS injury will

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not arrive at a hospital or other medical facility until some time after the injury, some degree of cell death will have occurred by the time the patient receives therapy. Thus, clinically relevant therapies require an appropriate therapeutic window such that they would afford neuroprotective effects when administered several hours to days following the insult. In addition, it is important to note that the therapeutic window for a particular therapy may vary with the specifics of the injury (type of insult, severity, etc), so additional research is needed to establish the therapeutic strategy that best fits each type of injury. For example, Hara et al. (1997) occluded the middle cerebral artery of adult rats for 2 hr and found that zVAD‐fmk was ineffective against cell loss when administered 1 hr after reperfusion. In contrast, Endres et al. (1998) occluded the middle cerebral artery for only 30 min and found that zVAD‐fmk could be delayed for 6 hr after reperfusion and still reduce infarct volume by 50% (similar to pretreatment). In some cases, combinatorial strategies may be required. Ma et al. (1998) found synergistic effects when mice were treated with both a caspase inhibitor (zVAD‐fmk or zDEVD‐fmk) and an excitotoxic inhibitor (MK‐801) prior to, or during, an ischemic insult. Not only was there greater neuroprotective potential with the combination treatment, but also the therapeutic window was wider than with either treatment alone. Zhao et al. (2004) recently demonstrated that 3 hr of mild hypothermia that began after a transient ischemic insult extended the therapeutic window for gene therapy by inhibiting cytochrome c release. Under those conditions, viral vectors that overexpressed Bcl‐2 were delivered as much as 5 hr after the insult and still improved the number of striatal neurons that survived to 2 days postinjury. Although treatments that reduce caspase activity and improve cell survival following injury are promising, the true test of any therapy is the effectiveness on behavioral recovery. One cannot assume that tissue (cellular) sparing will translate directly into behavioral sparing (recovery). First, cells that survive an injury may not be functioning normally, and there may be a threshold of cell survival that is required before functional improvements are observed (assuming that behavioral assessments are performed with an appropriately sensitive test). Second, the time course of neuroprotection is not necessarily the same as the time course for behavioral recovery. This principle was documented by Phillips et al. (2001) who used gene therapy to deliver calbindin to the hippocampus of rats after excitotoxic injury. Rats administered the vector either immediately or 1 hr after insult had equivalent improvements in neuron survival relative to controls at 2 days postinsult. In contrast, the performance of a hippocampus‐dependent spatial memory task was still disrupted at 7 days postinsult for rats that received the vector after a 1 hr delay and did not return to normal levels until postinjury day 11. Nevertheless, our continued understanding of (1) the events contributing to apoptotic cell death, (2) the activation and regulation of intracellular machinery involved, and (3) the development of methods to effectively reduce injury‐induced apoptosis will be essential for determining the clinical relevance of this cell death process in traumatic CNS injury.

Acknowledgments Preparation of this chapter was supported, in part, by a grant from the Kentucky Spinal Cord and Head Injury Research Trust, the National Institute for Neurological Diseases and Disorders, and the Cardinal Hill Research Endowment. While we attempted to include the most representative and descriptive studies, the enormous volume of literature in this field made it unfeasible to cite every study in this review. As such, the authors would like to apologize to our colleagues for any unintentional oversight or omission of their studies.

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18

Microglial Proteases

H. Nakanishi

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 552

2 2.1 2.2 2.3

Cathepsins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 553 Exogenous Antigen Presentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 553 Degradation of Extracellular Matrix Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 553 Intracellular Degradation of Ab Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 555

3 3.1 3.2 3.3

Plasminogen Activator, Plasmin, and Thrombin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 556 Degradation of ECM Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 556 Enhancement of NMDA Receptor‐Mediated Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 557 Activation of Microglia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 557

4

Proteasomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 558

5 Matrix Metalloproteases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 558 5.1 Degradation of ECM Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 558 5.2 Extracellular Degradation of Ab Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 559 6

Calpains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 559

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Caspases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 561

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Microglial proteases

Abstract: There is growing evidence that the proteolytic machinery of microglia is closely associated with their protective and cytotoxic roles in the central nervous system (CNS). Endosomal and lysosomal proteases including cathepsins E and S have been shown to play important roles in the major histocompatibility complex (MHC) class II‐mediated antigen presentation of microglia by processing of exogenous antigens and degradation of the invariant chain (Ii) associated with MHC class II molecules, respectively. There is evidence that some members of cathepsins are involved in extracellular proteolysis in addition to their functions in the endosomal–lysosomal system. We have recently found, by utilizing cathepsin S‐deficient mice, that cathepsin S is required for migration of microglia toward axotomized facial motoneurons. Several studies have suggested an involvement of cathepsin D in the clearance of amyloid‐b (Ab) peptides by microglia. On the other hand, attention has been also paid to deleterious effects of proteases secreted from microglia. Cathepsins S and B secreted from microglia are also involved in tissue damage and neuronal death. Moreover, tissue‐type plasminogen activator (tPA), a serine protease, secreted from microglia also participates in neuronal death, enhances N‐methyl‐D‐aspartate (NMDA) receptor‐mediated neuronal responses, and activates microglia via either its proteolytic or nonproteolytic activity. Calpain, a calcium‐dependent cysteine protease, has been demonstrated to play a pivotal role in the pathogenesis of multiple sclerosis (MS) by degrading myelin proteins extracellularly. Furthermore, matrix metalloproteases (MMPs) secreted from microglia also receive great attention as mediators of inflammation and tissue degradation through processing of proinflammatory cytokines and damage to the blood–brain barrier. Therefore, the accumulating knowledge about proteolytic events mediated by microglial proteases will contribute to better understanding of microglial functions in the CNS. List of Abbreviations: CLIP, class II-associated Ii peptide; CNS, central nervous system; ECM, extracellular matrix; L-LTP, late-phase LTP; LPS, lipopolysaccharide; LRP, lipoprotein receptor-related protein; MHC, major histocompatibility complex; MMPs, matrix metalloproteases; MS, multiple sclerosis; MS, multiple sclerosis; NMDA, N-methyl-D-aspartate; OVA, ovalbumin; PKC, protein kinase C; IFN-γ, interferon-γ; MAPK, mitogen-activated protein kinase; TIMPs, tissue inhibitors of metalloproteases; tPA, tissue-type plasminogen activator; uPA, Urokinase-type plasminogen activator

1

Introduction

Microglia are distributed in the central nervous system (CNS) and comprise up to 20% of the total glial cell population. In the normal CNS, microglia are present as ramified cells that have small cell bodies with numerous branching processes. In response to neuronal injury, ramified microglia rapidly transform into activated states. As an intermediate form, microglia have large cell bodies with several thicker processes. If neurons are severely injured, microglia further transform into phagocytic cells. Although activated microglia are known to have both beneficial and harmful effects, little is known about the molecular basis that determines the outcome of microglial activation in the CNS. There is growing evidence that microglia release proteases to affect neuronal functions (Nakanishi, 2003a–c, 2005). Protease activities are tightly regulated by at least three mechanisms: the gene transcription, the activation process of precursor forms, and the interaction with endogenous protease inhibitors. Once proteases are activated, they can exert irreversible cleavage of peptide bonds of various proteins. After cleavage, some substrates are inactivated and others are activated to gain new functions. Therefore microglial proteases are also considered to exert both beneficial and harmful effects. For better understanding of microglial functions, it is important to elucidate the key substrates for each microglial protease. Furthermore, better understanding the function of microglial proteases could contribute to the development of proteases and protease inhibitors as novel neuroprotective agents. In this chapter, I will focus on the proteolytic systems that are directly or indirectly associated with microglial functions in the CNS.


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Cathepsins

2.1 Exogenous Antigen Presentation A group of proteases in the endosomal–lysosomal proteolytic system is designated as cathepsins, from the Greek term meaning ‘‘to digest’’ (Nakanishi, 2003c). Some members of cathepsins are responsible for proteolytic pathways required for the major histocompatibility complex (MHC) class II‐mediated antigen presentation: a degradation of invariant chain (Ii) associated with MHC class II and endocytosed exogenous antigens. Although cysteine proteases including cathepsins S, B, L, and F are critical for the terminal step of Ii breakdown, their exact roles vary among different antigen‐presenting cells (Villadangos et al., 1999; Rieses and Chapman, 2000). In B cells and dendritic cells, cathepsin S exclusively mediates the degradation of Ii to class II‐associated Ii peptide (CLIP) during maturation of MHC class II molecules. For this purpose, thymic cortical cells and macrophages use cathepsin L and cathepsin F, respectively. In microglia, which are known to interact with infiltrated CD4þ T helper cells in the CNS, cathepsins S and L were suggested to be responsible for the degradation of Ii to CLIP because microglia express these proteases, but not cathepsin F (Gresser et al., 2001; Santambrogio et al., 2001). However, we used specific inhibitors for cathepsins to demonstrate that microglia utilize cathepsin S and/or cathepsin B but not cathepsin L in the degradation of Ii to CLIP (Nishioku et al., 2002). For the generation of antigenic peptides from endocytosed exogenous antigens, aspartic proteases including cathepsins E and D have been suggested in some antigen‐presenting cells, because aspartic protease inhibitors such as pepstatin A prevented antigen processing and Ii degradation (Maric et al., 1994). In B cells, expression level of cathepsin E was upregulated upon cellular activation and specific inhibitor of cathepsin E blocked the presentation of ovalbumin (OVA) (Bennett et al., 1992; Sealy et al., 1996). Furthermore, it has been reported that cathepsin D generated antigenic peptides from OVA or hen egg lysozyme that could be presented to T helper cells (Diment, 1990; Rodriguez and Diment, 1992; van Noort and Jacobs, 1994). More recently, however, experiments conducted with splenocytes and macrophages prepared from cathepsin D‐deficient mice have demonstrated that cathepsin D is dispensable for the degradation of Ii and for processing a number of exogenous antigens in splenocytes (Villadangos et al., 1997; Deussing et al., 1998). In microglia, we have demonstrated that cathepsin E is closely involved in the generation of antigenic peptides from OVA that could be presented to T helper cells based on the following observations: (1) cathepsin E is localized especially in endosome‐like structures of microglia, (2) pepstatin A, a potent hexapeptide inhibitor specific for aspartic proteases, significantly inhibited the interleukin‐ 2 production from OVA266–281‐specific T helper cell hybridomas upon stimulation with native OVA but not with OVA266–281 peptide (> Figure 18-1), (3) pepstatin A failed to block the degradation of Ii chain, (4) and microglia isolated from cathepsin D‐deficient mice retained the ability for antigen presentation (Sastradipura et al., 1998; Nishioku et al., 2002). The role of cathepsin E in antigen presentation by microglia has been further supported by the recent observation that cathepsin E is negatively regulated by class II transactivator, which is required for the expression of MHC class II and other genes related to antigen presentation (Yee et al., 2004). Although the precise implication of exogenous antigen presentation in the CNS is not fully understood, inflammatory cytokines secreted from activated helper T cells and microglia activated through their interaction may contribute to tissue damage and repair in autoimmune disease, viral infections, and chronic inflammatory disease.

2.2 Degradation of Extracellular Matrix Proteins There is substantial evidence that some cathepsins, especially cathepsins B and S, are involved in extracellular proteolysis in addition to their functions in the endosomal–lysosomal system. Cathepsin B has been

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. Figure 18-1 Effects of protease inhibitors on antigen presentation of naive ovalbumin (OVA) and antigenic peptide by primary cultured murine microglia. Microglia were isolated from a mixed primary culture prepared from 3‐day‐ old C57BL mice. After 10–14 days in culture, floating cells and weakly attached cells on the mixed primary cultured cell layer were isolated by gentle shaking of the flask. The resulting cell suspension was transferred to noncoated Petri dishes and allowed to adhere at 37 C. Unattached cells were removed after 30 min; microglia were isolated as strongly adhering cells. Each protease inhibitor (10 mM) was applied to culture medium 24 h before adding OVA‐specific helper T cell hybridomas and native OVA (1 mM) or OVA (266–281) peptide (0.3 mM). Supernatants from triplicate cultures were harvested, and helper T cell‐derived IL‐2 was measured by ELISA. Each column and bar represents mean SD of three experiments. The amount of IL‐2 in the absence of inhibitors was designated as control. Each value is expressed as a percentage of the amount to normalize the values with respect to control. Asterisks indicate significant differences from the control (**P < 0.01, ***P < 0.001 by Student’s t‐test)

suggested to participate in extracellular proteolysis. Ryan et al. (1995) used immortalized murine microglial cell line, BV‐2 cells, to demonstrate that cathepsin B was secreted as the heavy‐chain form, in addition to the proform, upon stimulation with lipopolysaccharide (LPS). Recently, it has been demonstrated that secreted cathepsin B from microglia is a major causative factor of microglia‐induced neuronal apoptosis (Kingham and Pocock, 2000, 2001). More recently, Gan et al. (2004) have conducted functional genomic approaches to identify cathepsin B as one of the genes transcriptionally induced by Ab peptides in BV‐2 cells. They have further shown that an inhibition of cathepsin B in BV‐2 cells using either small interference RNA‐ mediated gene silencing or a specific inhibitor for cathepsin B, CA074, leads to diminished the neurotoxic effects caused by Ab‐activated BV‐2 cells. These observations suggest an essential role for secreted cathepsin B from activated microglia in neuronal death mediated by degradation of extracellular matrix (ECM) proteins. Petanceska et al. (1996) have reported that there was a significant increase in cathepsin S activity secreted from both macrophages and microglia in response to LPS. They have also demonstrated that cathepsin S could degrade ECM proteins, including fibronectin, laminin, and proteoglycans even at neutral


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pH. Among lysosomal cysteine proteases, cathepsin S exceptionally retains a relatively wide pH optimum ranging from 5.0 to 7.5. Furthermore, Sukhova et al. (2003) have demonstrated that cathepsin S‐deficient monocytes and macrophages cannot migrate through artificial membrane containing smooth muscle cells, collagen mixture, and an endothelial monolayer. On the basis of these observations, it is likely that cathepsin S is closely associated with migration of monocytic cells by degradation of ECM proteins. Recently, we have found that the migratory activity of primary cultured microglia prepared from cathepsin S‐deficient mice was severely impaired. To further examine the role of cathepsin S, we have elucidated the role of cathepsin S in the migratory activity of microglia in cathepsin S‐deficient mice after axotomy of the facial motoneurons. Upon transection of facial nerves at the stylomastoid foramen, microglia accumulated in the axotomized side and spread on the surface of the axotomized facial motoneurons (> Figure 18-2a and > c). By contrast, cathepsin S‐deficient microglia touched on injured motor neurons, but they were unable to spread on the neuronal surface (> Figure 18-2b and > d). Furthermore, the mean number of microglia accumulated in the axotomized side of cathepsin S‐deficient mice was significantly smaller than that of the wild‐type mice (> Figure 18-2e). These observations indicate that cathepsin S is required for microglia to migrate and spread on the surface of axotomized facial motoneurons.

2.3 Intracellular Degradation of Ab Peptides It has been demonstrated that cathepsin D is responsible for the intracellular clearance of Ab peptides in human and rat brains (Hamazaki, 1996; McDermott and Gibson, 1996). Ab peptides are taken up . Figure 18-2 Microglial attachment to axotomized facial motoneurons 4 days after nerve transection. (a), contralateral and (c), axotomized sides of facial motor nuclei in wild‐type mouse. (b), contralateral and (d), axotomized sides of facial motor nuclei in cathepsin S‐deficient mouse. Brain sections including facial motor nuclei were immunostained for microglia using anti-lba1 antibody. In wild‐type mouse, microglia spread on the surface of the axotomized neurons. In cathepsin S‐deficient mouse, however, microglia touched on the axotomized neurons but were unable to spread on the surface. Scale bar ¼ 10mm. (e), the number of Iba1‐positive cells accumulated in the axotomized side of facial motor nuclei in wild‐type and cathepsin S‐deficient mice. Asterisk indicates a significant difference from the wild type (*P < 0.05 by Student’s t‐test)

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predominantly by microglia through class A scavenger receptors and class B scavenger receptor type I (Paresce et al., 1996; Husemann, 2001). Then, Ab peptides are accumulated and degraded in the endosomal– lysosomal systems of microglia (Paresce et al., 1997). It has been also reported that the degradation of Ab peptides in microglia is significantly inhibited by pepstatin A (Kakimura et al., 2002). These observations strongly suggest that phagocytosed Ab peptides are mainly degraded by cathepsin D in microglia. It is also noteworthy that immunization with Ab peptides has been demonstrated to reduce Ab peptides in transgenic mice with Ab plaques (Schenk et al., 1999). Anti‐Ab antibodies probably facilitate clearance of Ab peptides by driving microglia to phagocytose Ab peptides through Fc receptors. Thus, phagocytosis and subsequent degradation of Ab peptides by microglia may play a pivotal role in a strategy for the immunotherapy of Alzheimer’s disease.

3

Plasminogen Activator, Plasmin, and Thrombin

3.1 Degradation of ECM Proteins Tissue‐type plasminogen activator (tPA) is known to catalyze the conversion of plasminogen into plasmin, which plays an important role in fibrinolysis. In the CNS, tPA is distributed in discrete regions of the hippocampus, the hypothalamus, the amygdala, and the meningeal blood vessels (Nakanishi, 2005). Flavin et al. (2000) have demonstrated that secreted tPA is a principal factor for the activated microglia‐induced neuronal apoptosis because PA inhibitor‐1 (PAI‐1), tPA STOP, or coincubation with anti‐tPA antibody blocked it. More recently, however, Flavin and Zhao (2001) have reported that recombinant human tPA completely protected oxygen/glucose deprivation‐induced cell death of primary cultured rat hippocampal neurons. This protective effect of tPA was abolished by anti‐tPA antibody but not by PAI‐1 indicating that the protective effect of tPA is not mediated by proteolytic activity. Urokinase‐type plasminogen activator (uPA) secreted from microglia has been implicated in extracellular proteolysis, which is required for migration (Nakajima et al., 1992). When uPA binds to its surface receptor (uPAR), uPA activates plasminogen to plasmin, which, in turn, degrades ECM proteins such as laminin and fibronectin. Plasmin is also capable of activating the proenzyme of MMPs that are also known to degrade components of ECM. Strickland’s group has conducted a series of experiments to elucidate the mechanism underlying tPA‐ mediated excitotoxin‐induced degeneration in the hippocampus. They have first shown that mice deficient for tPA or plasminogen are resistant to seizure and neuronal death in the hippocampal CA1–CA3 subfields induced by excitotoxins (Tsirka et al., 1995, 1997). They also examined effects of intrahippocampal injection of protease inhibitors on kainate‐induced neuronal degeneration in the hippocampus of the wild‐type mice (Tsirka et al., 1995, 1997). When tPA inhibitor‐1 or a2‐antiplasmin was applied just prior to kainate administration, kainate‐induced neuronal death was markedly attenuated. These observations strongly suggest that plasmin formed by tPA‐mediated activation of plasminogen promotes neurodegeneration through degradation of ECM proteins. Finally, Chen and Strickland (1997) have demonstrated that laminin is the target ECM protein for plasmin and its degradation is responsible for excitotoxic neuronal death. Although hippocampal CA1 pyramidal cells are extremely sensitive to tPA‐mediated excitotoxin‐induced degeneration, the existence of tPA synthetic cells in the CA1 subfield is still controversial. Tsirka et al. (1995, 1997) combined a variety of approaches to show that tPA is synthesized by both hippocampal neurons and the satellite microglia that overlie the neuronal cell layer. More recently, Siao et al. (2003) have generated mice that exhibit restricted expression of tPA through introduction of tPA transgenes under the control of neuronal‐ or microglial‐specific promoters into tPA‐deficient mice. Using these animals, they observed that tPA, initially secreted from injured neurons, activates microglia at the site of injury. These activated microglia secrete additional tPA, which promotes degradation of ECM proteins leading to neurodegeneration by a plasminogen‐dependent mechanism. However, Salle´s and Strickland (2002) have reported that the CA1 subfield is essentially devoid of detectable tPA protein and activity. Furthermore, they could not detect tPA in the CA1 subfield even after excitotoxic injury. They have proposed several possible reasons why CA1 pyramidal cells that are devoid of tPA are most sensitive to tPA‐mediated excitotoxin‐induced neurodegeneration: (1) CA1 pyramidal cells may express an undetectable level of tPA that is crucial for neuronal


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degeneration; (2) neuronal death is likely to be attributable to a confluence of many factors; (3) and the presence of mossy fiber tPA may be required for neuronal death of CA1 pyramidal cells, possibly through feedback potentiation of electrical activity.

3.2 Enhancement of NMDA Receptor‐Mediated Responses There is increasing evidence that serine proteases, which are well known for their roles in blood coagulation and fibrinolysis, also contribute to potentiation of N‐methyl‐D‐aspartate (NMDA) receptor‐mediated responses in the CNS (Gingrich et al., 2000; Nicole et al., 2001). Gingrich et al. (2000) have demonstrated that the thrombin‐induced enhancement of NMDA receptor‐mediated responses is caused by proteolytic activation of the protease‐activated receptor‐1 (PAR‐1) but not by cleavage of NMDA receptors. On the other hand, tPA has been demonstrated to be necessary for late‐phase LTP (L‐LTP) in both the Schaffer collateral‐CA1 and mossy fiber‐CA3 pathways in the hippocampus by utilizing tPA‐deficient mice (Huang et al., 1996) and specific inhibitors for tPA (Baranes et al., 1998). Furthermore, mice overexpression of tPA showed an enhancement of LTP (Madani et al., 1999), and application of tPA also enhanced L‐LTP in rat hippocampal slices (Baranes et al., 1998). These observations strongly suggest that tPA plays a pivotal role in L‐LTP. Plasminogen, a major substrate of tPA, is converted to the broad‐spectrum protease plasmin, which can cleave ECM proteins such as laminin and fibronectin. However, plasmin‐ mediated extracellular proteolysis is unlikely to be a causative factor of tPA‐induced potentiation of L‐LTP, because plasmin impaired the maintenance of LTP by degrading laminin (Nakagami et al., 2000). Zhuo et al. (2000) have demonstrated that binding of tPA to its cell surface receptor, the low‐ density lipoprotein receptor‐related protein (LRP), activates cAMP‐dependent protein kinase A that plays a key role in L‐LTP. More recently, Nicole et al. (2001) have demonstrated that tPA enhances NMDA receptor‐mediated signaling by cleaving the NR1 subunit of the NMDA receptor. tPA was found to remove a fragment of approximately 15–20 kDa from the amino terminus of the NR1 subunit. They used coimmunoprecipitation experiments to show an apparent direct association between tPA and the NR1 subunit of the NMDA receptor in membrane preparations from cortical neuronal cultures. The cleaved form of the NR1 was also detected by coimmunoprecipitation assays after exposure of cortical neurons to NMDA. However, as pointed out by Matys and Strickland (2003), possible involvement of plasminogen and its activated form, plasmin, in the degradation of the NR1 subunit cannot be totally ruled out.

3.3 Activation of Microglia Tsirka and her colleagues have investigated the mechanism underlying the failure of microglial activation in tPA‐deficient mice during excitotoxic neuronal death (Tsirka et al., 1995, 1997). Rogove et al. (1999) have found that infusion of catalytically inactive tPA into the hippocampus of tPA‐deficient mice restored microglial activation, but failed to prevent neuronal death after injection of kainate suggesting that tPA mediates microglial activation through some mechanisms other than the conversion of plasminogen into plasmin. More recently, Siao and Tsirka (2002) extended their finding by demonstrating that tPA mediates microglial activation through its finger domain, which most likely interacts with annexin II, a cell‐surface receptor, to initiate an intracellular signaling cascade. Thus, tPA appears to associate with excitotoxic neuronal death through two pathways (Pawlak and Strickland, 2002; Tsirka, 2002). The first is the proteolytic pathway for the activation of plasminogen to plasmin that degrades plasmin, leading to neuronal death. The second is the nonproteolytic pathway for the activation of microglia that contribute to neurodegeneration through the release of neurotoxic molecules after cellular activation. Thrombin is also known to activate various cell types, including neurons, astrocytes, and microglia. The thrombin‐induced activation of neurons and astrocytes is mediated by the proteolytically cleaving PAR‐1. Proteolytic activation of PAR‐1 results in sequential activation of tyrosine kinase and RhoA, a member of the Ras family of small GTP‐binding proteins. Activated RhoA is translocated to the plasma membrane

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where it can transduce the signal to its effector proteins, leading to cellular activation and apoptosis. In microglia, thrombin induces NO release and iNOS expression (Mo¨ller et al., 2000; Ryu et al., 2000). These effects, however, may be not mediated by proteolytic cleavage of PAR‐1 because a peptide agonist could not mimic the effect of thrombin. Thrombin‐induced NO release and iNOS expression were significantly suppressed by inhibitors of protein kinase C (PKC), mitogen‐activated protein kinase (MAPK), or nuclear factor kB (NF‐kB) indicating that thrombin activates microglia via PKC, MAPK, and NF‐kB (Ryu et al., 2000). By contrast, Suo et al. (2002) have demonstrated an essential role of PAR‐1 in the thrombin‐induced microglial activation by utilizing PAR‐1‐deficient primary cultured microglia. More recently, however, Hanisch et al. (2004) have demonstrated that neither PAR activation nor proteolytic‐ or ligand‐like activities of thrombin proper were essential for microglial activation.

4

Proteasomes

A proteasome is a large multiunit complex (26S proteasome) that composes a 20S core particle, the catalytic component, and two 19S regulatory complexes that are attached to either end of the catalytic core. The proteasome degrades polyubiquitinated proteins to small peptides. The MHC class I antigen presentation pathway requires the 20S proteasome and two types of activator proteins, PA28 and PA700, to generate antigenic peptides. The antigenic peptides are transported by a transporter associated with antigen processing into the endoplasmic reticulum, where they become associated with MHC class I molecules and are then translocated to the cell surface to activate cytotoxic T lymphocytes. In response to interferon‐g (IFN‐g), three constitutive b subunits of the 20S proteasome are replaced by the immunoproteasome subunits. IFN‐g‐ induced subunit replacement is implicated in increasing the epitope capacity of the 20S proteasome. It has been suggested that microglia are involved in MHC class I‐mediated antigen presentation during viral infection (Kolson et al., 1998). Recently, Stohwasser et al. (2000) characterized the dynamics of the 20S–26S proteasome subunit composition of primary cultured murine microglia and BV‐2 cells in response to IFN‐g and LPS. Both IFN‐g and LPS induced immunoproteasomes in primary cells, whereas BV‐2 cells responded only to IFN‐g. These observations support the idea that microglia play a pivotal role in MHC class I‐mediated antigen presentation during viral and bacterial infections in the CNS.

5

Matrix Metalloproteases

5.1 Degradation of ECM Proteins MMPs are initially synthesized as the precursor form with Zn2þ of the catalytic domain bound to the cysteine residue of the propeptide region. Various factors can activate proMMPs through a conformational change that disrupts the cysteine–Zn2þ binding (cysteine switch) and leads to the expression of the catalytic site. The intermediate form of MMPs is converted to the mature enzyme through autocatalytic cleavage of the prodomain. In the CNS, MMP‐2 (gelatinase A) and MMP‐9 (gelatinase B) are secreted by microglia and astrocytes as active forms (Colton et al., 1993; Chauvet et al., 2001). Their proteolytic activities in the extracellular space are regulated by tissue inhibitors of metalloproteases (TIMPs), natural inhibitors of MMPs, secreted from microglia and other cells (Cross et al., 1999). It has been recently suggested that MMP‐2 and MMP‐9 contribute to cerebrovascular diseases by degrading components of the basal lamina around the cerebral blood vessels such as type IV collagen, laminin, and fibronectin. In fact, MMP inhibitors or MMP‐neutralizing antibodies reduce edema and infarction in animal models of stroke (Romanic et al., 1998; Rosenberg et al., 1998; Asahi et al., 2000; Jiang et al., 2001). Furthermore, a broad‐spectrum MMP inhibitor, BB94, can reduce kainate‐induced neuronal cell death and leukocyte recruitment in the striatum (Campbell et al., 2004). In recent studies utilizing gene‐targeted mice, MMP‐9 has been further suggested to be a critical mediator for brain injuries, because deficiency of MMP‐9, but not MMP‐2, results in the reduction of lesion volumes caused by traumatic brain injury, focal ischemia (Lo et al., 2002), or spinal cord injury (Noble at al., 2002).


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5.2 Extracellular Degradation of Ab Peptides Recently, Selkoe’s group has screened certain neuronal and nonneuronal cell lines to ascertain whether they constitutively release proteases capable of degrading Ab (Qiu et al., 1997). Among the cell lines examined, they found that BV‐2 produces the most potent Ab‐degrading activity, which was completely blocked by an MMP inhibitor. After purification and characterization of Ab‐degrading protease secreted from microglia, they have concluded that it is an insulin‐degrading enzyme, a thiol metalloendopeptidase, that degrades small peptides such as insulin, glucagons, and atrial natriuretic peptide (Qiu et al., 1998). Insulin‐degrading enzyme was also found to be capable of mediating the oligomerization of Ab.

6

Calpains

Calpains are a family of Ca2þ‐dependent cysteine proteases, expressed as a precursor form that undergoes autocatalytic processing to yield the mature form in a Ca2þ‐dependent mechanism (in micromolar and millimolar concentrations for m‐calpain and m‐calpain, respectively). The level of calpain activity is regulated by interaction with calpastatin, a specific endogenous inhibitor. In multiple sclerosis (MS), an autoimmune demyelination disease, the degradation of myelin proteins results in the destabilization of myelin sheath. Calpains have been implicated in myelinolysis because myelin proteins including myelin . Figure 18-3 Activation of m‐calpain in microglia after treatment with ATP or 20 ‐30 ‐O‐(benzoyl‐benzoyl) ATP (BzATP). Microglia were isolated from a mixed primary culture prepared from 3‐day‐old Wistar rats. Immunoblots from SDS‐ polyacrylamide gel electrophoresis of extracts from microglia treated with ATP (1mM) or BzATP (100 mM) were developed with antiactive m‐calpain IgG. Rabbit polyclonal antibody against the synthetic peptide LGRHE, which matches the N‐terminal sequence of the active form of human m‐calpain, was used as the primary antibody. Each column and bar represents mean SD of three experiments. Each value is expressed as a percentage of the density to normalize the value with respect to the density of the control protein band. Asterisks indicate significant differences from the control (**P < 0.01, ***P < 0.001 by Student’s t‐test)

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basic proteins are their substrates, and calpains are significantly increased in the white matter from the brains of patients with MS (Shields et al., 1999). Further, calpains were increasingly expressed in activated microglia, macrophages, and mononuclear phagocytes in the white matter from the brains of patients with MS (Shields et al., 1998). On the basis of these observations, Shields et al. have hypothesized that extracellular calpains secreted from glial cells and inflammatory cells may degrade myelin proteins to produce immunologic fragments engulfed by antigen‐processing cells for MHC class II antigen presentation. In the white matter of the aged rhesus monkey, m‐calpain derived from microglia has been also demonstrated to be involved in myelin protein metabolism (Hinman et al., 2004). Although calpain activation in microglia has been implicated in the pathology of the demyelinating diseases, little is known about the mechanism of calpain activation in microglia. Recently, we have found that m‐calpain in microglia was activated upon stimulation with ATP. As shown in > Figure 18-3, the 76‐kDa isoform of the catalytic subunit of m‐calpain corresponding to the active form in primary cultured microglia was significantly increased following addition of ATP or 20 ‐30 ‐O‐(benzoyl‐benzoyl) ATP (BzATP), a selective P2X7 receptor agonist. It was also noted that the amount of active m‐calpain was significantly reduced at 60 min after treatment with ATP. This may be due to auto‐degradation or secretion of m‐calpain after activation. Extracellular ATP signaling through P2X7 receptors can directly associate with m‐calpain activation because these receptors work as a Ca2þ‐permeable ligand‐gated channel (Inoue, 1998). Therefore, it is reasonable to consider that ATP leaked from damaged cells is involved in m‐calpain activation through P2X7 receptors in microglia accumulated in pathological sites.

7

Caspases

Caspases, designated as cysteine‐dependent aspartate‐specific proteases, are a family of proteases that cleave after aspartate residues in their substrates. Among the caspase family, a great deal of data has shown that . Figure 18-4 Microglial protease functions. Ab, amyloid‐b; ECM, extracellular matrix; Ii, invariant chain; MBP, myelin basic protein; MMP‐2, matrix metalloprotease‐2; MMP‐9, matrix metalloprotease‐9; NMDA, N‐methyl‐D‐aspartate; tPA, tissue‐plasminogen activator; uPA, urokinase‐type plasminogen activator


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caspase‐3 activation plays a central role in apoptosis, cleaving various important cytoplasmic and nuclear proteins. We have reported that relatively low concentrations of neurotoxins such as 6‐hydroxydopamine and methylmercury induce apoptosis of primary cultured rat microglia mainly through activation of caspase‐3‐like proteases and partially through endosomal/lysosomal proteases (Takai et al., 1998; Nishioku et al., 2000). The apoptosis of microglia induced by 6‐hydroxydopamine and methylmercury compromises the host defense system and homeostasis of the CNS and may have an important pathogenic implication in Parkinson’s disease and Minamata disease, respectively. Ferrari et al. (1999) have demonstrated that ATP simultaneously initiates both caspase‐3‐dependent apoptotic and necrotic pathways in microglia through P2X7 receptors because caspase‐3 inhibitors completely prevented ATP‐induced appearance of nuclear apoptotic morphology such as chromatin condensation and margination but did not affect necrotic alterations such as cell swelling and vacuolization of the cytoplasm. Furthermore, it has been also reported that an overstimulation of microglia activated caspases to induce apoptosis (Kingham and Pocock, 2000; Lee et al., 2001; Liu et al., 2001). These observations strongly suggest that overactivation‐induced apoptosis of microglia is an auto‐regulatory mechanism that downregulates the number of activated microglia after termination of the pathological stimuli in the CNS.

8

Conclusion

Although the levels of the transcripts or the endogenous inhibitors of proteases in the CNS strictly regulate their activities, both intracellular and secretory proteases of microglia are closely associated with both beneficial and harmful roles of microglia in the CNS. > Figure 18-4 summarizes both the intracellular and extracellular proteolytic cascade of microglial proteases. The growing understanding of proteolytic systems that are directly or indirectly associated with microglial functions could also contribute to the development of pharmacological interventions for various disorders in the CNS.

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Microglial proteases Nishioku T, Takai N, Miyamaoto K–I, Murao K, Hara C, et al. 2000. Involvement of caspase 3‐like protease in methylmercury‐induced apoptosis of primary cultured rat cerebral microglia. Brain Res 871: 160-164. Noble LJ, Donovan F, Igarashi T, Goussev S, Werb Z. 2002. Matrix metalloproteinases limit functional recovery after spinal cord injury by modulation of early vascular events. J Neurosci 22: 7526-7535. Paresce DM, Chung H, Maxfield FR. 1997. Slow degradation of aggregates of the Alzheimer’s disease amyloid b‐protein by microglial cells. J Biol Chem 46: 29390-29397. Paresce DM, Ghosh RN, Maxfield FR. 1996. Microglial cells internalize aggregates of the Alzheimer’s disease amyloid b‐protein via scavenger receptor. Neuron 17: 553-565. Pawlak R, Strickland S. 2002. Tissue plasminogen activator and seizures: A clot‐buster’s secret life. J Clin Invest 109: 1529-1531. Petanceska S, Canoll P, Devi LAL. 1996. Expression of rat cathepsin S in phagocytic cells. J Biol Chem 271: 4403-4409. Qiu WQ, Walsh DMW, Ye Z, Vekrellis K, Zhang J, et al. 1998. Insulin‐degrading enzyme regulates extracellular levels of amyloid b‐protein by degradation. J Biol Chem 273: 32730-32738. Qiu WQ, Ye Z, Kholodenko D, Seubert P, Selkoe DJ. 1997. Degradation of amyloid‐b protein by a metalloprotease secreted by microglia and other neural and nonneural cells. J Biol Chem 272: 6641-6646. Riese RJ, Chapman HA. 2000. Cathepsins and compartmentalization in antigen presentation. Curr Opin Immunol 12: 107-113. Rodriguez GM, Diment S. 1992. Role of cathepsin D in antigen presentation of ovalbumin. J Immunol 19: 2894-2898. Rogove A, Saio C‐J, Keyt B, Strickland S, Tsirka, SE. 1999. Activation of microglia reveals a nonproteolytic cytokine function tissue plasminogen activator in the central nervous system. J Cell Sci 112: 4007-4016. Romanic AM, White RF, Arleth AJ, Ohlstein EH, Barone, FC. 1998. Matrix metalloprotease expression increases after cerebral focal ischemia in rats. Stroke 29: 1020-1030. Rosenberg GA, Estrada EY, Dencoff JE. 1998. Matrix metalloprotease and TIMPs are associated with blood–brain barrier opening after reperfusion in rat brain. Stroke 29: 2189-2195. Ryan RE, Sloane BF, Sameni M, Wood PL. 1995. Microglial cathepsin B: An immunological examination of cellular and secreted species. J Neurochem 65: 1035-1045. Ryu J, Ryo H, Jou I, Joe E. 2000. Thrombin induces NO release from cultured rst Microglia via protein kinase C,

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19

MMPs and Other Matrix‐Degrading Metalloproteinases in Neurological Disease

P. E. Gottschall . K. Conant

1

Extracellular Matrix in the CNS and Matrix‐Degrading Proteinases . . . . . . . . . . . . . . . . . . . . . . . . 566

2 2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.1.5 2.1.6 2.2 2.3 2.4

The Matrix Metalloproteinase and Related Families of ECM‐Degrading Proteinases . . . . . . . 567 Matrix Metalloproteinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 567 Collagenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 568 Gelatinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 568 Stromelysins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 Matrilysins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 Membrane‐Type MMPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 Other MMPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 A Disintegrin and Metalloproteinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 570 A Disintegrin and Metalloproteinase with Thrombospondin Motifs . . . . . . . . . . . . . . . . . . . . . . . . . . . 570 Inhibitors of the MMPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 572

3 3.1 3.2 3.3 3.4 3.4.1 3.4.2 3.4.3

Matrix Metalloproteinases in Neurological Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 572 Multiple Sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 574 Cerebral Ischemia and Opening of the Blood–Brain Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 575 Glioma Invasion and Angiogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 576 Alzheimer’s Disease and Other Neurodegenerative Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 578 Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 578 HIV Dementia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 579 Other Neurodegenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 581

4

Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 581

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MMPs and other matrix‐degrading metalloproteinases in neurological disease

Abstract: The metzincin superfamily of metalloproteinases includes the most well‐studied matrix metalloproteinases (MMPs), the ADAMs (a disintegrin and metalloproteinase), and the ADAMTSs (ADAM with thrombospondin repeats) families of extracellular matrix (ECM)‐degrading enzymes. These proteases are mostly secreted with important exceptions for the transmembrane sheddases and can degrade all protein components of the ECM. In addition to ECM proteins, secreted and cell‐surface proteins including growth factors, cytokines, and chemokines may be cleaved by the MMPs. These proteases alter the composition and structural organization of the ECM; and importantly, cleavage affects intracellular signaling induced by binding of matrix molecules to cell‐surface receptors. A body of data has been generated on the expression of these proteases in neurological disease, which is summarized here. In addition, the role of individual MMPs in particular animal models of disease has been investigated using genetically targeted mutant mice. In general, these results have suggested a detrimental role for the MMPs that relates to the induction of their expression in response to injury or disease. List of Abbreviations: ADAMs, a distintegrin and metalloproteinase; ADAMTS, ADAM with thrombospondin repeats; ECM, extracellular matrix; GPI, glycosylphosphatidylinositol; MT-MMPs, membrane-type MMPs; TIMPs, tissue inhibitors of metalloproteinases

1

Extracellular Matrix in the CNS and Matrix‐Degrading Proteinases

Extracellular matrix (ECM) in the neuropil of the central nervous system (CNS) is markedly different in its component molecules and aggregate matrix compared to other tissues. Most obviously, the matrix in the nervous system lacks fibrillar collagen that many other tissues depend upon for integrity of structure. The CNS ECM is high in proteoglycan content, especially the aggregating proteoglycans that bind to hyaluronic acid. These proteoglycans belong to the lectican family of aggregating, chondroitin sulfate‐substituted proteoglycans and include brevican, versican, aggrecan, and neurocan. The N‐terminal globular domain of the lecticans binds to hyaluronic acid whereas the C terminus binds to tenascin (Aspberg et al., 1997; Kleene and Schachner, 2004), and along with ‘‘link protein’’, these molecules create a three‐dimensional lattice which is thought to stabilize cellular structures within nervous tissue (Hockfield et al., 1990). These aggregates are often found surrounding neurons as ‘‘perineuronal nets’’ but are also deposited in neuropil and are even found in white matter (Yamaguchi, 2000; Gottschall and Conant, unpublished observations). The lattice complex is similar to that found in cartilage with the major exception that collagen is not present, and the abundance of the glycosaminoglycan side chains is much lower in brain compared to cartilage. Since these polysaccharide side chains can become hydrated, their lower concentration in brain results in a matrix that occupies a much smaller relative volume compared to cartilage. However, the cerebrovasculature, unlike the neuropil, is surrounded by a basal lamina that is similar if not identical to that found in other tissues, with the exception that astrocytes, along with endothelia, contribute to its formation (Nag, 2003). Interaction between the ECM and cell‐surface molecules on neurons and glia provide fundamental signals during development and in regeneration after injury, and also have a role in synaptic plasticity in the adult (Dityatev and Schachner, 2003; Kleene and Schachner, 2004). These protein interactions at the cell surface provide information to the neuron or glial cell about the extracellular microenvironment, and the cells respond accordingly. A good example of such a phenomenon is the transmembrane proteoglycan syndecan‐ 2 that is enriched in dendritic spines of developing neurons, and when overexpressed and clustered on the spine membrane, it induces spine morphogenesis and maturation (Ethell and Yamaguchi, 1999; Ethell et al., 2001). This action is dependent upon its intracellular PDZ‐binding domain and intraspine signaling events that affect rearrangements in synaptic scaffold proteins and the neuronal cytoskeleton (Ethell et al., 2001). There are a number of ECM proteins that avidly bind to the heparan sulfate (and chondroitin sulfate) chains that are covalently linked to the ectodomain of syndecans. Thus, interactions between syndecan‐2, heparan‐ binding, and ECM molecules present around the synaptic cleft may modulate spine morphogenesis. Because matrix–neural and matrix–glial interactions are pivotal in nervous system development and maintenance, any mechanism that leads to changes in the ECM content or conformation would also be of critical import. A set of diverse proteinases cleave, degrade, and turnover the ECM in the brain.


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The metzincin superfamily of metalloproteinases includes the most well‐studied matrix metalloproteinases (MMPs), the ADAMs (a disintegrin and metalloproteinase), and the ADAMTSs (ADAM with thrombospondin repeats). The MMPs are mostly secreted proteases (with important exceptions, see below); they have the broadest substrate specificity among these families, and can degrade all protein components of the ECM. In addition to ECM proteins, secreted and cell‐surface proteins including growth factors, cytokines, and chemokines may be cleaved by the MMPs (Sternlicht and Werb, 2001), and methods have been developed to rapidly scan for novel substrates of the MMPs (Overall et al., 2002). In fact, the key role of MMP‐2 in dampening the inflammatory action of monocyte chemoattractant protein‐3 was discovered using these methods (McQuibban et al., 2000). The ADAM family is structurally similar to the MMPs except that they are nearly all transmembrane proteins and contain a disintegrin domain that may or may not bear an RGD, integrin‐binding, cell adhesion sequence. Most of the ADAMs act mainly as adhesive proteins as only several of these molecules have been shown to be proteolytically active. However, ADAMs that are active are efficient as ectodomain ‘‘sheddases,’’ proteinases that cleave the extracellular domains of transmembrane or glycosylphosphatidylinositol (GPI)‐anchored proteins that include several ‘‘receptors’’. In the example of spine maturation described above, the action of a syndecan‐cleaving ‘‘sheddase,’’ which is known to be active (Bernfield et al., 1999), would modulate spine morphogenesis and maturation by increasing or decreasing its activity to regulate the abundance of syndecan‐2 ectodomain on the cell surface, and thus, the traffic of signaling that is important for the maturation of spines. The ADAMTSs are somewhat special because they are secreted proteases that are deposited in and bind to the ECM. Each of the proteolytically active ADAMTSs exhibits a reasonably selective activity but among them their activity is diverse and ranges from proteoglycanase to procollagen N‐proteinase to von Willebrand factor proteinase activity. The additional domains present in the ADAMs and ADAMTSs such as the disintegrin and thrombospondin repeats likely act to localize these proteins for interaction with binding partners. Although much more is known about the structure and especially the function of MMPs in physiology and pathophysiology, it will likely turn out that the ADAMs and the ADAMTSs will be as important as the MMPs in disease states. For instance, it was recently demonstrated that ADAMTS5 is the major aggrecan‐ degrading proteinase in cartilage (Stanton et al., 2005) and that a deficiency of ADAMTS5 alone prevents cartilage degradation in a mouse model of osteoarthritis (Glasson et al., 2005).

2

The Matrix Metalloproteinase and Related Families of ECM‐Degrading Proteinases

The MMPs, ADAMs, and ADAMTSs are the most important metalloproteinases involved in ECM turnover. The serine proteases, tissue plasminogen activator, and urokinase‐type plasminogen activator as well as plasmin are also capable of degrading ECM proteins, but their description is beyond the scope of this chapter.

2.1 Matrix Metalloproteinases The MMPs are a 24‐member (23 in humans) family of ECM‐degrading proteinases, which are expressed as secreted, type I transmembrane (MMP‐14, MMP‐15, MMP‐16, and MMP‐24) or GPI‐anchored (MMP‐17 and MMP‐25) proteins. All are Caþþ‐ and Znþþ‐dependent proteinases, which are synthesized as inactive zymogens with common structure. A signal peptide is expressed at the N terminus to direct the protein to the secretory pathway, followed by a propeptide region that is required to confer latency to the protease, and a catalytic domain that binds Znþþ in a triple‐histidine‐binding pocket. A hinge region of variable length occurs next, which usually links the catalytic domain with the C‐terminal hemopexin domain, a region important for substrate binding and interaction with the endogenous tissue inhibitors of metalloproteinases (TIMPs) (Sternlicht and Werb, 2001; Visse and Nagase, 2003). In addition to Znþþ in the active site, the signature for proteases in this family is a conserved sequence in the propeptide region, ‘‘‐PCRGXPD‐,’’ called the ‘‘cysteine switch’’ motif. This unpaired cysteine residue interacts with Znþþ in the active site which excludes water and maintains the MMP in its zymogen form. Disruption of this interaction by

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organomercurials or denaturants results in a conformational change; water replaces the thiol group resulting in activation of the zymogen, autocatalytic cleavage of the propeptide, and full activation of the protease. In vivo, there is evidence that the MMPs are cleaved by other active proteases resulting in proteolytic activation in a cascade sort of manner, although the concept is not firmly established. Other evidence suggests that in conditions of inflammation, reactive oxygen or nitrogen species are effective at disrupting the ‘‘cysteine–zinc’’ interaction, and thus have the potential to activate MMPs. Due to their potential for substantial tissue destruction, the expression and activation of MMPs are tightly regulated. Regulation of MMP activity may occur by modulation of gene transcription and translation, by changing the mechanisms of proteolytic (or other) activation of the MMP from its latent state, or by altering inhibitor interaction with the MMP. Regulation of gene expression of the MMPs is a major controlling mechanism for most MMPs and is complex (the complexity illustrated in an excellent, lengthy review of MMP‐9 activity; Van den Steen et al. (2002)). Generally, however, increased expression is observed during periods of tissue pathology (e.g., inflammation), modeling, or remodeling (e.g., during development or after injury) or even during physiological periods of tissue turnover in the adult (e.g., ovulation). Cytokines, chemokines, growth factors, oncogenes, matrix proteins, adhesion molecules, matrix protein fragments, ultraviolet radiation, and hormones (steroid and others) are key regulators of MMP gene expression (Nagase and Woessner, 1999; Sternlicht and Werb, 2001). Corticosteroids inhibit cytokine‐stimulated increases in MMP gene transcription in vitro, and the dogma states that during periods of inflammation cytokine‐induced increases in MMP expression may be limited by elevated levels of glucocorticoids. Cleavage of the propeptide domains of the MMPs by other proteases, in a cascade‐like manner, may result in their activation, as supported by in vitro evidence (Nagase, 1997); however, animal models have been used to confirm the concept (Carmeliet et al., 1997). An interesting complex interplay among the MMPs that requires MMP‐14 (MT1‐MMP) and TIMP‐2 for localization of MMP‐2 at the cell surface is apparently the mechanism by which MMP‐2 is activated (Murphy et al., 2003), although it is puzzling that the phenotype of the MMP‐14 deficient mouse bears no resemblance to those mice deficient in TIMP‐2 (Holmbeck et al., 2004) or MMP‐2. In addition, as described in more detail below, several of the MMPs bear proprotein convertase consensus processing sites, with the ADAMs and the ADAMTSs, that allow for intracellular cleavage of the enzymes by furin‐like proteases (Pei and Weiss, 1995). The individual MMPs may be subdivided into several categories based on common structure and substrate specificity. For a complete, detailed review of the MMPs and their substrates, see McCawley and Matrisian (2001) and Sternlicht and Werb (2001).

2.1.1 Collagenases The ‘‘collagenases’’ (MMP‐1, MMP‐13, and MMP‐18) are identified by their unique ability to cleave fibrillary collagen types I, II, and III at a single site about three‐quarters of the way down the sequence from the N terminus. In addition, these proteases are efficient in the cleavage of chondroitin sulfate proteoglycans and a number of other ECM molecules as well as other substrates. In humans, MMP‐1 immunoreactivity appears to be abundant in white matter microglia (Dickson et al., 1996). MMP‐1 protein stimulates integrin‐mediated signaling involving dephosphorylation of the PI3/Akt survival pathway, and activates caspases and neuronal death in a manner independent of its proteolytic activity (Conant et al., 2004). A recent study has shown that MMP‐1 can also cleave and thus activate proteinase‐activated receptor‐1 (PAR‐1) on breast cancer cells (Boire et al., 2005). This may be significant in that PAR‐1 may be expressed on neurons and its activation by thrombin has been linked to cell death (Dery et al., 1998; Turgeon et al., 1998; Festoff et al., 2000; Hollenberg, 2003; Vergnolle et al., 2003).

2.1.2 Gelatinases MMP‐2 and MMP‐9 constitute the gelatinases; they bear the common names of gelatinase A (72 kDa) and gelatinase B (92 kDa), respectively, and both proteases are highly effective at degrading type IV collagen and


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denatured collagen or gelatin in addition to other important substrates. Each gelatinase has three tandem repeats of a type II fibronectin module, inserted following the catalytic domain, which bind laminin, collagens, and gelatin and affect substrate specificity of the protease (Allan et al., 1995). More information is available about these MMPs in the nervous system than any others, likely because pro‐MMP‐2 is abundantly expressed by several cell types in brain, especially in astrocytic end‐feet surrounding blood vessels (Rosenberg, 2002) and in other astroglia (Rivera et al., 2002), and there are simple and direct methods to quantify levels of pro‐MMP‐2 and pro‐MMP‐9 (and their active forms) (Zhang and Gottschall, 1997; Snoek‐van Beurden and Von den Hoff, 2005). MMP‐9 is detected in select populations of neurons, notably hippocampal pyramidal neurons in human sections (Backstrom et al., 1996) and in pyramidal and dentate granule neurons in rats (Rivera et al., 2002). It is highly expressed by activated microglia and reactive astrocytes and especially by invading leukocytes during periods of inflammation (Yong et al., 2001) and after ischemic injury (Rivera et al., 2002)

2.1.3 Stromelysins MMP‐3 (stromelysin 1) and MMP‐10 (stromelysin 2) have broad, yet similar substrate specificities (Sternlicht and Werb, 2001; Visse and Nagase, 2003) and include the full spectrum of ECM proteins. Exposure of the aggregating proteoglycan, aggrecan, to MMP‐3 results in the cleavage of an Asn‐Phe bond (IPEN * FFGV) and in the production of fragments similar to those found in human cartilage in vivo. MMP‐3 is effective at cleaving nervous system proteoglycans as well, and along with the ADAMTSs may be important for the turnover of these proteoglycans in brain (Nakamura et al., 2000). A recent study has also suggested that MMP‐3 may play a role in microglial cell activation by showing that the active, but not the latent, proform of MMP‐3 markedly stimulated the expression of proinflammatory cytokines in cultured microglia (Kim et al., 2005). Interestingly, conditioned media derived from MMP‐3‐activated microglia turned out to have potent neurotoxic activity.

2.1.4 Matrilysins Matrilysins (MMP‐7 and MMP‐26) are identified by the lack of a C‐terminal hemopexin domain. Like the stromelysins, MMP‐7 has a broad substrate specificity yet has been associated with a diverse number of epithelial‐derived tumors (Wielockx et al., 2004) and has a role in the ectodomain shedding of several key transmembrane proteins such as pro‐a‐defensin, pro‐tumor necrosis factor‐a (TNF‐a), and Fas ligand (McCawley and Matrisian, 2001).

2.1.5 Membrane‐Type MMPs Of the six so‐called ‘‘membrane‐type MMPs’’ (MT‐MMPs), four are type I membrane proteins (MMP‐14, MMP‐15, MMP‐16, MMP‐24) while the remaining two are anchored to the membrane via a GPI linkage (MMP‐17 and MMP‐25). ECM substrates cleaved by the MT‐MMPs seem to be as broad as those for the secreted MMPs but in addition they are crucially involved in activation mechanisms of the MMPs, especially in the activation of MMP‐2 (Visse and Nagase, 2003). In the nervous system, MMP‐24 is widely and selectively expressed in the brain with the highest expression in the cerebellum, hippocampal pyramidal and dentate granule neurons, and in neurons of the brain stem (Sekine‐Aizawa et al., 2001).

2.1.6 Other MMPs The most well studied of the uncategorized MMPs in the nervous system is MMP‐12, or metalloelastase, a protease that appears to regulate oligodendrocyte maturation and morphological differentiation (Larsen

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and Yong, 2004) but has a complex role after injury in the nervous system. MMP‐12 appears to be mainly expressed by macrophages outside the nervous system and modulates macrophage migration (Shipley et al., 1996). MMP‐12 can cleave a number of ECM proteins including elastin.

2.2 A Disintegrin and Metalloproteinase The ADAM family of proteins (about 30 members in humans) has been shown to be involved in biological functions related to adhesion and proteolytic processing that range from development to reproductive sperm–egg fusion to angiogenesis (for review, see White (2003); Blobel (2005); White Web site of table of ADAMs: http://www.people.virginia.edu/jw7g/Table_of_the_ADAMs.html). The two most evident differences between the MMPs and proteins of the ADAM family are: the ADAM proteins contain a disintegrin domain and they are integral type I membrane proteins. The prototype ADAM consists of several characteristic domains that are conserved among individual members of the family. The N‐terminal signal sequence and prodomain are proceeded by a metalloproteinase catalytic domain, the disintegrin domain, a cysteine‐rich sequence, an epidermal growth factor‐like domain, and a transmembrane and cytoplasmic domain. About half of the 30 ADAM genes contain the HEXXH motif for metalloproteinases, providing at least the potential for catalytic activity, and ADAMs 9, 10, 12, 15, 17, 19, 28, and 33 have been shown to cleave protein substrates (Blobel, 2005). In the nervous system, the ADAMs are best known for their crucial roles in protein ectodomain shedding and have been shown to have key biological roles ranging from the mediation of Notch signaling (Mumm and Kopan, 2000) to the regulation of axonal growth (Galko and Tesseir‐Lavigne, 2000) and contact‐mediated inhibition of axon growth (Hattori et al., 2000). In addition, in particular circumstances, the ADAMs modulate amyloid precursor protein (APP) processing (Allinson et al., 2003), an important action related to Alzheimer’s disease (AD) (see below). Moreover, ADAMs alter the biological action of various cytokines and growth factors as well as their receptors that are ‘‘shed’’ in the nervous system (for review please see Huovila et al. (2005) and White (2003)). Of note, several ADAMs are highly expressed in the developing and adult nervous systems (Weskamp et al., 2002; Kieseier et al., 2003; Toft‐Hansen et al., 2004).

2.3 A Disintegrin and Metalloproteinase with Thrombospondin Motifs The ADAMTSs are another large, 19‐member family of metalloproteinases that fall within the metzincin superfamily (for a complete review, see Apte (2004); Porter et al. (2005)). The first ADAMTS was cloned from a cachexigenic tumor cell line (Kuno et al., 1997) and a related molecule was later discovered to be highly effective in cleaving the matrix protein aggrecan, thus ending the search for the elusive, cartilage‐ derived ‘‘aggrecanase’’ activity (Tortorella et al., 1999). However, many of the ADAMTSs do not have a known substrate or are not proteolytically active. Those that are proteolytically active may be categorized into subgroups with distinct substrate specificities. For example, ADAMTS‐2, ‐3, and ‐14 exhibit procollagen N‐proteinase activity (Colige et al., 1997; Vazquez et al., 1999; Fernandes. et al., 2001), ADAMTS‐13 is a von Willebrand factor‐cleaving protease (Fujikawa et al., 2001) and mutations in this gene cause thrombotic thrombocytopenia purpura (Levy et al., 2001; Zheng et al., 2001), and ADAMTS‐1, ‐4, ‐5, ‐8, ‐9, and ‐15 are efficient aggrecanases and have activity on related proteoglycans (Tortorella et al., 1999; Sandy and Verscharen, 2001; Sandy et al., 2001; Rodriguez‐Mazaneque et al., 2002; Somerville et al., 2003; Collins‐ Racie et al., 2004). Like the MMPs and ADAMs, the ADAMTSs contain a signal peptide for secretion, a propeptide domain, and the Znþþ‐containing catalytic domain. The major structural differences between members of the ADAM family and the ADAMTS proteins are that the ADAMTSs contain (sequentially) a variable number (1–18) of thrombospondin repeats or modules that follow the cysteine‐rich region, a spacer domain, and sometimes a C‐terminal module (Apte, 2004; see Apte Web site of ADAMTSs: http://www.lerner.ccf.org/bme/apte/adamts/). Although it is unclear whether the propeptide domain of the


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ADAMTSs is essential for conferring latency to the proteases, each of the ADAMTSs bears a proprotein convertase consensus sequence (RXR/KR) that is cleaved by furin or furin‐like convertases in the Golgi apparatus during processing of the protein. C‐terminal cleavage is important for substrate specificity and further activation of the ADAMTSs. Truncation of the C‐terminal end either by autocatalyic cleavage or cleavage by other proteases (likely at the cell surface) in the spacer domain results in the appearance of 53 and 40 kDa isoforms of ADAMTS4 from the 68 kDa isoform with the C terminus intact (Flannery et al., 2003; Gao et al., 2004). Although the shorter isoforms may show reduced affinity for aggrecan, they may be more proteolytically active (Gao et al., 2002). An important functional distinction between the ADAMTSs and most of the MMPs is that many of the ADAMTSs bind to the ECM near or at the cell surface, a property probably due to the presence of the thrombospondin repeats. The importance of the ADAMTSs in the nervous system is reflected by the abundance and appearance of fragments of brevican (Matthews et al., 2000; Yuan et al., 2002), versican (Westling et al., 2004), and aggrecan (Yuan et al., 2002) with the ADAMTS’ signature glutamyl endopeptidase neoepitope (Yamaguchi, 2000), and conserved ADAMTS‐specific cleavage sites have been identified in each of these proteoglycans (> Figure 19-1). ADAMTS1 and ADAMTS4 are reasonably abundant; they are expressed in hippocampal pyramidal neurons and dentate granule neurons and expression of both appear to be upregulated after excitotoxic injury to the brain (Yuan et al., 2002). The localization of the ADAMTS‐derived neoepitope of

. Figure 19-1 Illustration of human brevican glutamyl endopeptidase cleavage site for the ADAMTSs (top). Cleavage of human brevican by ADAMTS proteinase results in the generation of a short N‐terminal fragment of brevican with a ‘‘neoepitope’’ C‐terminal sequence ‘‘‐EATESE‐.’’ This fragment may be selectively identified with a specific antibody on Western blot or in immunohistochemistry. The human, mouse, rat, and bovine ADAMTS cleavage sites in versican V2 and aggrecan are shown (bottom)

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these proteoglycans is associated with increased expression of the ADAMTSs in vivo (see below). ADAMTS activity is negatively regulated by TGFb in astrocytes in vitro (Hamel et al., 2005). The capacity for multisite cleavage of brevican by the ADAMTSs and the MMPs suggests a complex regulation of brevican turnover and fragment formation (Nakamura et al., 2000).

2.4 Inhibitors of the MMPs Because many of the proteases of the metzincin family would uncontrollably cleave their substrates, a regulatory mechanism exists for dampening this activity in vivo in the form of endogenous inhibitors termed TIMPs. TIMPs are expressed early in phylogeny from invertebrates to mammals, and in mammals four members of this family have been identified. The four TIMPs vary in molecular weight from 21 to 36 kDa and each contains six conserved cysteine residues that form three disulfide bonds to maintain conformation of the N‐terminal domain with the C‐terminal end of the molecule. TIMP‐1 is the most highly regulated of the TIMPs and increases its expression in response to injury or inflammation often in tandem with several MMPs, especially MMP‐9 (Rivera et al., 1997; Pagenstecher et al., 1998). In adult CNS, TIMP‐1 is expressed at very low levels (as opposed to reasonable expression in other tissues); TIMP‐2 and TIMP‐3 are found in neurons and astrocytes in the CNS although the distribution between them differs markedly. TIMP‐2 is highly expressed in cerebral cortex, cerebellum, brain stem, and spinal motor neurons (Pagenstecher et al., 1998), whereas TIMP‐3 is found in thalamus, subventricular zone, and olfactory bulb (Jaworski and Fager, 2000). Although among the TIMPs, TIMP‐4 is the molecule most selectively expressed in the nervous system, little is known about its role. In addition, the TIMPS exhibit actions independent of their ability to inhibit metalloproteinase activity. Most recently TIMP‐2 has been found to stimulate neurite outgrowth and promote neuronal differentiation in vitro by acting as an antimitogenic signal in an MMP‐ independent fashion. TIMP‐2 appears to signal via interaction with a3b1 integrin that activates the cAMP/ Rap1/ERK pathway. Such a finding has significant implications for the ability of the adult brain to generate new neurons after injury or with disease (Perez‐Martinez and Jaworski, 2005). Another protein that may inhibit MMP activity is the membrane‐anchored glycoprotein RECK (reversion‐inducing cysteine‐rich protein with Kazal motifs). This glycoprotein has been shown to inhibit MMP‐2, MMP‐9, and MT1‐MMP (Oh et al., 2001) activity. Mice null for RECK die at embryonic day 10.5 with defects in vascular development and collagen fibrils (Oh et al., 2001). This phenotype can be partially rescued by an MMP‐2 null mutation. Of interest, the ability of RECK to inhibit tumor invasion and metastasis may be suppressed in Epstein‐Barr virus (EBV) disease, in that EBV latent‐membrane protein‐1 targets RECK (Liu et al., 2003).

3

Matrix Metalloproteinases in Neurological Disease

In disorders of the nervous system, the condition that includes the secretion of MMPs and the expression of other molecules in response to pathology is referred to as ‘‘inflammation.’’ Depending on the disease or injury, this term ‘‘inflammation’’ is used to describe a wide‐range and diverse set of conditions that may exist in the nervous system. The condition of ‘‘inflammation’’ may be derived from the innate and/or acquired immune response, or it may be the result of invasion of peripheral leukocytes into the brain, or it may be caused by cells intrinsic to the nervous system. For instance, in multiple sclerosis (MS), a well‐ known characteristic of the pathology is the infiltration of leukocytes (lymphocytes and macrophages) from the peripheral blood into the brain parenchyma, an action thought to be a component of an autoimmune response. These invading leukocytes contribute to extracellular levels of MMPs; however, these cell types are not the only source of MMPs and other proinflammatory molecules present. Associated with an MS lesion are activated astrocytes and reactive microglia, in addition to neurons that are demyelinating or may be undergoing axonal injury, and along with endothelial cells, these cells also express MMPs in the disease state. In contrast to MS, ‘‘inflammation’’ associated with AD pathology lacks significant numbers of T cells, neutrophils, or other cells types from the systemic circulation (with the possible exception of monocytes/


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macrophages which may be recruited from the circulation), but there are activated astrocytes that surround neuritic and senile plaques, reactive microglia embedded within the plaques, and more rarely activated perivascular microglia. There is good evidence that these cells types found intrinsic to the brain are the predominant types that contribute to inflammation in AD. Thus, when the term ‘‘inflammation’’ is used, it relates to the proinflammatory nature in the context of a particular disease, not to any rigidly defined individual state or condition. Finally, it should be noted that the function of altered MMP expression in neurological disorders that is associated with inflammation may not be entirely detrimental, and may have remodeling/regenerative actions. In most of these diseases, proinflammatory cytokines produced by reactive microglia are thought to act on astrocytes to induce expression of MMPs. Reactive microglia themselves likely contribute a different set of MMPs as do neurons that respond to the ‘‘inflammatory’’ conditions. Although cytokines are found in diseased tissues, there is quite a variation that occurs among the diseases regarding the profile of MMPs that are found (Yong et al., 2001). However, with the glial cell types in culture, it is clear how they respond. Astrocytes derived from neonatal rats produce MMP‐2 (Apodaca et al., 1990) and treatment with lipopolysaccharide (LPS), interleukin‐1b (IL‐1b), TNF‐a, or phorbol esters induce the secretion of MMP‐9 and MMP‐3 (Gottschall and Yu, 1995; Gottschall and Deb, 1996; Witek‐Zawada and Koj, 2003) via a mechanism that involves MAP kinase signaling in the case of phorbol (Arai et al., 2003) and IL‐1b (Wu et al., 2004). Compared to MMP‐3 and MMP‐9, smaller increases in MMP‐10, MMP‐12, and MMP‐13 gene expression were observed after LPS treatment of rat astrocytes (Wells et al., 1996). Rat microglia treated with LPS, lectins, or other activators increase the production of the pro‐ and active forms of MMP‐9 and MMP‐3 (Gottschall et al., 1995; Gottschall and Deb, 1996; Rosenberg et al., 2001). The chemokines, MCP1, MIP1b, RANTES, IL‐8, and fractalkine were found to increase the secretion of MMPs and TIMPs from a human fetal microglial cell line and several of these cytokines were also effective in inducing MMP expression in rat brain microglia (Cross and Woodroofe, 1999). Thus, these in vitro results should be placed in context with the changes observed in MMP expression in brain tissue extracts or when MMPs are localized using immunohistochemistry in acute and chronic neurological disease. In addition, it is difficult to determine the actual activational state of these metalloproteinases in vivo or in vitro due to their zymogen nature and to identify a substrate for a particular MMP whose expression is changed in some disease, usually chronic disease. Using immunohistochemistry, the metalloproteinase may be localized, and it may even be possible to raise an antibody that will only recognize the active isoform of the protease; however, even so, the metalloproteinase may be secreted into an environment that contains abundant levels of an inhibitor of the protease. Biochemical techniques to measure the MMPs have similar obstacles—lower molecular weight forms of a protease identified on a Western blot or zymogram may indeed be an active form, but may be secreted in tandem with the endogenous inhibitor, usually a TIMP. Thus, often an investigation will attempt to estimate the balance of active metalloproteinase to the levels of inhibitor present and how this ratio changes during treatment or disease. One attempt at circumventing this problem is to identify a substrate for the metalloproteinase and measure levels of the specific cleavage fragment using antibodies that will only recognize the fragment and not the parent protein substrate. The real advantage of such a technique is that an active protease can be identified and localized in brain tissue sections. The cleavage sites in a number of ECM (and other protein) substrates have been identified, and usually these are metalloproteinase family‐specific cleavage sites (ADAMTS cleavage sites for brevican, versican, and aggrecan are shown in > Figure 19-1). Antibodies may be raised against the neoepitope sequences that are generated by cleavage. The technique of using neoepitope‐specific antibodies to recognize protease‐derived fragments of matrix proteins has been used in cartilage research for many years (Sandy et al., 1991). Of course, when levels of these fragments are measured, it is difficult if not impossible to identify which specific member of the particular proteinase family was responsible for the generation of the fragment from the substrate, although the expression of a particular protease in that region where the cleaved fragment is located certainly provides strong evidence. Thus, when interpreting data that measures MMP (or other metalloproteinase) levels or location in diseased tissues, it is important to view this data from the perspective that the MMP may not be an active MMP, and that, even if it happens to be a cleaved isoform, TIMP levels determine whether it is active or not.

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3.1 Multiple Sclerosis MS is a disease of the white matter whose course may be progressive or relapsing–remitting and the severity of the disease may vary from a benign disease that has no impact on patient lifestyle to a rapidly evolving and incapacitating disease (Kaspar, 2005). MS lesions are characterized by perivenular cuffing with an infiltrate into the surrounding white matter consisting predominantly of T cells and macrophages, and together with the response from endogenous neural cells, the infiltrate regions are areas with elevated concentrations of proinflammatory cytokines and chemotactic chemokines, many of which act to induce the production of MMPs (Nelissen et al., 2003). In regions of inflammation, the blood–brain barrier is breached, which may be related to the action of MMPs (Rosenberg et al., 1996b). In most cases, myelin‐ reactive T cells are present, and in more than half the cases autoantibodies are produced against myelin that promote demyelination and stimulate macrophages and microglia to scavenge the myelin debris. As a lesion evolves, there may be some differentiation of preoligodendrocytes that attempt to remyelinate surviving naked axons. In addition, astrocytes become active as characterized by an increase in glial fibrillary acidic protein expression. Although some MS lesions may be axon sparing, evidence suggests that axon loss is the major cause of the irreversible neurological disability in MS (Kaspar, 2005). Major therapeutics developed to treat MS have targeted the immune system responses that accompany the exacerbations of the disease. One of the newer and most effective treatments has been interferon (IFN)‐b, and in some cases, the antiimmune action of IFN‐b is successful to mitigate the course of the disease. Since there are mononuclear cells derived from the bloodstream present in the lesions of MS, it follows that migration of these cells from the blood through tissues requires a catabolic environment, and the MMPs are good candidates to mediate this migratory action. The MMPs would be involved in the breakdown of the vascular basement membrane and other ECM proteins, allowing for chemotactic migration of these mononuclear cells from the peripheral blood. In the 1990s, it was observed that MMP‐9 was elevated in cerebrospinal fluid (CSF) from patients with MS and other inflammatory neurological disease (Gijbels et al., 1992), and the concept that there are elevated levels of MMP‐9 secreted from these mononuclear cells in and around lesions in MS is now firmly established (Opdenakker and Van Damme, 1994; Cuzner et al., 1996; Cossins et al., 1997). Interestingly, MMP‐9 was the only MMP of several measured that was significantly elevated in relapsing–remitting disease irrespective of the relapse or stable phase (Leppert et al., 1998). The levels and activity of MMP‐9 appeared to be related to cell count in the CSF (Yushchenko et al., 2000) and although microglia and macrophages contribute to its production, the Th1 subset of CD4þ T cells was shown to produce higher levels of MMP‐9 than do Th2 cells (Abraham et al., 2005). Mainly, activated microglia associated with MS lesions and perivascular cuffs and activated astrocytes were MMP‐9 immunopositive (Cuzner et al., 1996; Maeda and Sobel, 1996; Cossins et al., 1997). Although the best case is made for elevations in MMP‐9 in and around MS lesions, other MMPs, including MMP‐2, MMP‐3, MMP‐7, and MMP12 (Maeda and Sobel, 1996; Anthony et al., 1997; Cossins et al., 1997; Veldhuis et al., 2003) are elevated in MS. Because TNF‐a is a cytokine thought to be involved in the pathogenesis of MS including stimulating the release of MMPs, it is interesting that the protease, ADAM17, responsible for cleaving it into the mature form is elevated in MS and expressed by invading T cells in the parenchyma (Kieseier et al., 2003). In addition, in a screen of mRNA for numerous metalloproteinases, ADAM12 was the only ADAM found to be elevated in tissue from patients with MS (Toft‐Hansen et al., 2004). There are a host of potential mechanisms as to how MMP‐9 might be involved in the MS disease process and these have been summarized in a thorough and intriguing review (Opdenakker et al., 2003). Because they provide a strong rationale for using MMP inhibitors as therapeutics agents, some of these will be discussed here. As outlined above, elevated levels of MMP‐9 are found in MS lesions and in CSF of MS patients, whereas the concentrations of the MMP inhibitors, the TIMPs (especially TIMP‐1), are reduced in MS (Lee et al., 1999), markedly elevating the MMP‐9/TIMP ratio, and this holds true even in peripheral mononuclear cells derived from MS patients (Lichtinghagen et al., 1999; Kouwenhoven et al., 2002). Two autoreactive antigens that have received attention in MS are myelin basic protein and aB‐crystallin. MMP‐9 has the ability to cleave each protein resulting in proteolytic ‘‘remnant epitopes,’’ which may be autoreactive and induce encephalitis (Proost et al., 1993; van Noort et al., 1995). Interestingly, the most effective available treatment for MS is IFN‐b and IFN‐b has been shown to inhibit MMP‐9 expression in glial cells


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(Liuzzi et al., 2004) and suppress T cell migration (Leppert et al., 1996). MMP‐9 levels were lower in CSF obtained from MS patients treated with IFN‐b (Jacobs et al., 1996). IFN‐b itself may be cleaved and inactivated by MMP‐9 (Nelissen et al., 2003) suggesting that combination therapy with IFN‐b and an MMP‐9 inhibitor may be able to elevate local IFN concentrations. It should be noted that many of the findings regarding alterations in MMP‐9 expression and activity also follow, and have been extended to the most well‐characterized animal model of the disease, the experimental autoimmune encephalitis (EAE) model. MMP‐9 is upregulated during the course of EAE and young mice deficient in MMP‐9 by specific gene deletion were resistant to the development of EAE. Moreover, inhibition of MMP activity using a broad spectrum small molecule inhibitor (Gijbels et al., 1994) or the antiinflammatory antibiotic minocycline (Brundula et al., 2002) protected animals against the harmful effects of EAE. Minocycline is a potent, direct MMP‐9 inhibitor (Paemen et al., 1996). Finally, statins, whose effects include inhibition of MMP expression, have shown promise in EAE and are being considered for use in MS (Greenwood et al., 2003; Stuve et al., 2003).

3.2 Cerebral Ischemia and Opening of the Blood–Brain Barrier Ischemia in the CNS can occur as a result of focal occlusion of a cerebral vessel, which causes a reduction in blood flow to the region of the brain that it innervates, or, as in the example of a cardiac arrest, a global cerebral ischemia results in the death of cells that are most susceptible to the hypoxic conditions. As a result, each of these conditions has a distinct pathology. With acute occlusion of an artery, a reduction in flow to zero results in early and substantive loss of brain tissue, whereas a modest reduction in flow which is restored over a short time may cause a patient to exhibit only transient symptoms, i.e., transient ischemic attack. Within the central core of a major infarct where the loss of flow is greatest, ionic disturbances, the release of excitotoxic agents, low ATP and energy stores, and metabolic failure due to the loss of flow cause severe and early damage that in most cases is irreversible (Lo et al., 2003). However, a significant proportion of the tissue damage due to stroke is delayed, and occurs from hours to days after the ischemic event in a zone, which is peripheral to the core, called the ischemic penumbra. Most agents developed over the last several years and tested preclinically for the treatment of stroke are targeted toward the protection of neurons that are lost in the penumbra during this delayed period. The advantage of such an agent compared to what is available today is that penumbra‐targeted agents may be administered at longer times after symptoms of the stroke have appeared. The mechanisms responsible for damage in penumbra after a focal ischemic episode are varied, numerous, and complex and are clearly beyond the scope of this chapter. For a comprehensive look at these details, see Lo et al. (2003). It is now well established that during a focal ischemic episode in the brain, there is an upregulation of multiple metalloproteinases; for review see Cunningham et al. (2005). Both MMP‐2 and MMP‐9 activities are increased after focal cerebral ischemia in human tissue (Clark et al., 1997). In rats undergoing middle cerebral artery occlusion, MMP‐9 expression and activity is rapidly induced at 3 h and at 12 and 24 h after transient focal ischemia in the rat, whereas MMP‐2 was not increased until 5 days after the infarct (Rosenberg et al., 1996a; Fujimura et al., 1999; Gasche et al., 1999). There were marked increases in active gelatinase activity observed in zones within the infarcted area in the basal ganglia (Loy et al., 2002) and some evidence suggests that MMP‐2 may be activated in addition to MMP‐9 (Chang et al., 2003). Leukocytes that invade the parenchyma secrete MMPs in the area of the infarct (Justicia et al., 2003), and inhibiting the migration of the infiltrating leukocytes limits the size of the infarct (Veldhuis et al., 2003). MMP‐3 is likely elevated in the infarct and contributes to the damage in addition to the gelatinases (Sole et al., 2004; Gu et al., 2005), although a neuroprotective role for MMP‐3 has also been suggested (Cunningham et al., 2005). TIMP‐1 mRNA expression is markedly elevated as well after transient ischemia (Wang et al., 1998; Rivera et al., 2002). Conceptually, the role of metalloproteinases after ischemia may be multifold and includes the loss of the basal lamina that disrupts the blood–brain barrier and causes the edema observed in the damaged region, a direct action on neurons resulting in apoptotic death, remodeling, and angiogenic effects for recovery. A plethora of evidence indicates that MMPs are partly responsible for at least the first two of these

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actions. The earliest demonstration of the involvement of the MMPs in disruption of the blood–brain barrier was that injection of MMP‐2 directly into brain parenchyma altered the ultrastructure of the basal lamina and allowed the infiltration of normally excluded substrates followed by extravasation of blood cells into the region. Each of these actions was blocked by a metalloproteinase inhibitor (Rosenberg et al., 1992). The size of the infarct induced by focal ischemia in an MMP‐9 deficient mouse (Asahi et al., 2000) or after administration of a small molecule MMP inhibitor (Romanic et al., 1998) was markedly smaller compared to the size in appropriate control mice, and there appeared to be evidence for reduced damage to the blood– brain barrier in the MMP‐9 deficient mouse (Asahi et al., 2001). It may be that MMP‐9 derived from leukocytes is most important for loss of integrity of the blood–brain barrier after transient ischemia (Gidday et al., 2005). The antiinflammatory antibiotic minocycline, which is known to directly inhibit MMP activity, reduces infarct size after focal ischemia, but only in the wild‐type and not in the MMP‐9 deficient mouse indicating that minocycline is exerting its protective effects by blocking the action of MMP‐9 (Koistinaho et al., 2005). These data suggest a major role for MMP‐9 in infarct development in animal models, and more recent results indicate this may hold for human stroke as well (Rosell et al., 2005). Significantly less is known about the potential remodeling and proangiogenic effects of the MMPs and in vivo models of cerebral ischemia; it would be difficult to dissect out these positive effects from the seemingly more substantive role(s) the MMPs play in the development of the infarct.

3.3 Glioma Invasion and Angiogenesis Gliomas are a type of human tumor that often have a poor prognosis and survival rate despite significant advances in medical and surgical oncology. Most adult gliomas are astrocytomas, which are the more aggressive grade III (anaplastic astrocytomas) or grade IV (glioblastoma multiforme) type. Clinically, glioblastoma multiforme may be classified into secondary glioblastomas that develop slowly by progression through stages of slower growing astrocytomas and later to an aggressive grade tumor, whereas primary glioblastomas develop extremely rapidly without radiological or morphological evidence of a lesser malignant precursor (Ohgaki, 2005). Patients with primary glioblastomas usually have a short clinical history ( Table 20-1) (Laskowski and Kato, 1980; McDonald, 1985). The structure, binding to proteases, and mode of activity of the inhibitors, have been extensively studied and are comprehensively reviewed elsewhere (Bode and Huber, 1992; Tumminello et al., 1993; Potempa et al., 1994; Roberts et al., 1995; Fumagalli et al., 1999; Hibbetts et al., 1999; Forsyth et al., 2001; Janciauskiene, 2001; Turk et al., 2001; Onda et al., 2005).

2

Nonspecific Protease Inhibitors: a‐Macroglobulins

The major function of a‐macroglobulins, a group of high‐molecular weight nonspecific protease inhibitors, is rapid inhibition of excess proteolytic activity due to either endogenous or exogenous proteases. Various . Table 20-1 Protease inhibitors Inhibitor class Nonspecific protease inhibitors Class‐specific protease inhibitors

a‐Macroglobulins Serine protease inhibitors

Cysteine protease inhibitors Metalloprotease inhibitors Aspartic protease inhibitors

Example a2‐Macroglobulin a1‐Antichymotrypsin, protease nexin I, protease nexin II, neuroserpin, plasminogen activator inhibitor, antithrombin III, pigment epithelium‐derived factor Cystatin A, cystatin B, cystatin C Tissue inhibitors of metalloproteases (TIMP‐1, ‐2, ‐3, ‐4) Pepstatin

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studies have shown the involvement of a‐macroglobulins in coagulation (Downing et al., 1978; Ellis et al., 1982), fibrinolytic systems (Gonias, 1992), inflammatory conditions such as pancreatitis (McMahon et al., 1984; Banks et al., 1991), sepsis (Abbink et al., 1991; de Boer et al., 1993), disseminated intravascular coagulation (Hellgren et al., 1984; Lammle et al., 1984), and in adult respiratory distress syndrome (Velasco et al., 1986). a2‐macroglobulin (a2‐M) is the most extensively studied nonspecific protease inhibitor belonging to this group of nonspecific protease inhibitors. It is a major serum glycoprotein composed of four identical subunits of approximately 185 kDa each (Chaudhuri, 1993). It is widely expressed by many tissues (Thomas et al., 1989; Gaddy‐Kurten and Richards, 1991), primarily in the liver (Borth, 1992; Chu and Pizzo, 1994), but also in adult and fetal brain (Dziegielewska et al., 1986) in human cortical and hippocampal neurons (Strauss et al., 1992) and is secreted into various body fluids, including cerebrospinal fluid (CSF) (Garton et al., 1991). The a2‐M receptor, low‐density lipoprotein‐related protein (LRP) clears a2‐M–protease complexes from serum and biological fluids via endocytosis (Kounnas et al., 1995). The idea that a2‐M is an essential inhibitor is underscored by the fact that it has been well preserved throughout evolution (Starkey and Barrett, 1982; Sottrup‐Jensen et al., 1990) and that no complete deficiencies of the inhibitor have been reported in humans (Sottrup‐Jensen, 1989). Homologous deletion of the receptor gene is lethal at embryonic stages and exhibits profound effects on brain development (Herz et al., 1992). These data indicate that the a2‐M–LRP system is essential for normal development.

2.1 a2‐M in the Nervous System and Neurological Disorders a2‐M binds noncovalently, yet with different specificities and to different degrees, with cytokines, including tumor necrosis factor (TNF)‐a, interleukin (IL)‐6, platelet‐derived growth factor, and neurotrophins such as nerve growth factor and neurotrophin‐3 (Liebl and Koo, 1994). As a carrier of cytokines, a2‐M is involved in the brain inflammatory response to injury, which involves recruitment of inflammatory cells and astrocytes that secrete growth factors/cytokines, among them IL‐6 (Akira et al., 1990; Perry et al., 1993), which in turn can stimulate cerebral neurons and astroglia to synthesize and secrete a2‐M (Ganter et al., 1991). Indeed, increased a2‐M immunoreactivity was detected in glial cells in human brain following tissue injury or neoplastic transformation (Lopes et al., 1994). A regulation of a2‐M activity involves the formation of a monoamine‐activated form of the inhibitor resulting in loss of the ability to inhibit proteases and enhanced binding to LRP (Imber and Pizzo, 1981). Activated a2‐M can reduce the levels of monoamines often associated with neurodegenerative diseases and aging (Kish et al., 1992) and can inhibit neurite outgrowth and survival of embryonic sensory and cerebral cortical neurons (Liebl and Koo, 1993). It was also shown that activated a2‐M inhibits choline acetyltransferase activity (Liebl, and Koo, 1994). Thus, a2‐M has multiple important functions under physiological conditions, and may be involved in the brain response to neuronal injury and age‐related neurodegenerative pathologies. It has been related to several diseases in the nervous system. A polymorphism of a2‐M is linked with argyrophilic grain disease, a neurodegenerative disorder of the aged human brain associated with the formation of abnormal tau protein in specific neurons and microglial cells (Ghebremedhin et al., 2002). The level of a2‐M increases 3.5‐fold in tick‐borne encephalitis patients and remains stable during the acute period of the disease (Merzeniuk et al., 2000). a2‐M is also involved in regulation of increased proteolytic activity occurring in multiple sclerosis (Jensen et al., 2004). Alzheimer’s disease. a2‐M has been implicated in AD by its detection in amyloid plaques, tangles, and dystrophic neurites in the brain, the major pathological characteristics of AD patients (Tooyama et al., 1993; Van Gool et al., 1993). a2‐M is a candidate gene for AD (Saunders et al., 2003) and for CAA (Yamada et al., 1999). Genetic linkage analyses have suggested the presence of an AD locus on chromosome 12, in the vicinity of the A2M gene (Rogaeva et al., 1998; Wu et al., 1998; Mayeux et al., 2002; Myers et al., 2002) or the LRP gene (Pericak‐Vance et al., 1997; Rogaeva et al., 1998; Scott et al., 1999; Scott et al., 2000). However, genetic association analyses for AD and A2M have been controversial. The discrepancy between the generally positive association findings in family‐based samples and the generally negative association

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findings in case‐control samples further suggest that A2M may be a risk factor primarily in individuals with a family history of AD (Saunders and Tanzi, 2003). High specific binding of a2‐M and Ab, the amyloid peptide deposited in the brain of AD patients, was shown, attenuating Ab fibril formation and associated neurotoxicity (Du et al., 1998; Hughes et al., 1998). Furthermore, protease‐activated a2‐M and Ab complexes can mediate Ab catabolism (Lauer et al., 2001) or can undergo LRP‐mediated endocytosis followed by trafficking of Ab to the lysosome for degradation or translocation out of the brain and into the plasma (Narita et al., 1997). However, while a2‐M is involved in Ab degradation and clearance, an excess of a2‐M can have neurotoxic effects (Kovacs, 2000; Shibata et al., 2000; Lauer et al., 2001; Friedland, 2002) possibly by promoting and sustaining inflammatory responses (McGeer and McGeer, 2001).

3

Class‐Specific Protease Inhibitors

The four class‐specific protease inhibitors have been grouped based on the type of proteases they inhibit. The most commonly used classification of proteases is based on their catalytic mechanism to cleave peptide bonds. The name of each class refers to the amino acid that either directly or indirectly, via polarization of a water molecule, attacks the carbonyl carbon of the scissile peptide bond in substrates (Broome and Petanceska, 2001). The protease inhibitors bind reversibly either directly to the active site of the protease or to surface sites adjacent to the active site, preventing access of substrates to the active site of the protease (Bode and Huber, 1992).

3.1 Serine Protease Inhibitors Members of the largest superfamily of class‐specific protease inhibitors inhibit proteases that contain serine residues in their active sites. These proteases play integral roles in physiologic processes, including digestion (trypsin), blood coagulation (coagulation factors), immune reactions (complement components and neutrophil elastase), and fibrinolysis and fertilization of the ovum (Gettins et al., 1993; van Gent et al., 2003). At least four distinct families of serine protease inhibitors exist in mammals: Kunitz, Kazal, leuko‐ proteases, and serpins. (1) Kunitz inhibitors bind their related enzymes in a substrate‐like manner (Bode and Huber, 1991). Members of this family contain inhibitory domains with six conserved cysteine residues that form three disulfide bonds, contributing to the compact structure of the proteins. They inhibit trypsin, chymotrypsin, plasmin, kallikrein, elastase, cathepsin G, and coagulation factors (Brown et al., 1978; Creighton and Charles, 1987). The amyloid precursor protein (APP) belongs to this family of inhibitors. (2) Similar to Kunitz inhibitors, the Kazal family members have three conserved disulfide bonds and a reactive site loop for each inhibitory domain. In vitro studies demonstrated that Kazal inhibitors could inhibit trypsin, chymotrypsin, elastase, and subtilisin (Laskowski and Kato, 1980). (3) The two inhibitory domains of leuko‐protease inhibitors contain four conserved disulfides each and the mechanism of inhibition is similar to that of Kunitz and Kazal inhibitors. They inhibit elastase and cathepsin G. (Stetler et al., 1986; Thompson and Ohlsson, 1986; Simmen et al., 1992; Alkemade et al., 1994). (4) The serpin family contains at least 60 members with sequence similarity, conserved amino acid residues, and tertiary structure. They differ in the degree of glycosylation and ability to dimerize. Novel proteins have been classified as serpin‐like based on their sequence (Hammond et al., 1987; Stein et al., 1989; Hammond, 1990; Potempa et al., 1994; Bird, 1998) such as endopin 1 and endopin 2 (Hwang et al., 1999; Hook and Hwang, 2002), and phosphatidylethanolamine‐binding protein (Hengst et al., 2001). Active and inactive forms of serpins are present in biological fluids. They are very abundant in mammalian plasma and play an important role in many physiologic processes, primarily to neutralize overexpressed serine proteinase activity (Travis and Salvesen, 1983). Alterations in functional levels of a serpin by a change in structure and/or secretion may have pathological implications. Serpin dysfunction in the brain has been implicated in neurodegenerative disorders, mental disorders, and familial encephalopathy.


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Serine protease inhibitors in the nervous system and neurological disorders. Serine proteases and their serpin inhibitors have been detected in neurons and in glial cells of the central nervous system (CNS). Secretion of the proteases regulates the balance of accumulation and degradation of the extracellular matrix. The proteases and their serpin inhibitors were found at sites of neuronal injury and it was suggested that serpin levels affect the outcome of brain injury (Smith‐Swintosky et al., 1995; Buisson et al., 1998). Serine proteases and their inhibitors are extensively involved in inflammatory processes related to neurodegenerative diseases. APP (protease nexin II). APP is an integral membrane protein, which contains a large ectodomain secreted by proteolytic cleavage of the protein at sites close to the plasma membrane. The APP gene consists of 19 exons, located on chromosome 21, which are alternatively spliced into several mRNA forms, two of which, encoding 751‐ and 770 amino acids, contain a Kunitz‐type inhibitory domain (Goldgaber et al., 1987; Kang et al., 1987; Robakis et al., 1987; Tanzi et al., 1987; Kitaguchi et al., 1988). In the brain, APP695 isoform is predominantly found in neurons while APP751 and APP770 isoforms are predominantly found in astroglial cells (Rosa et al., 2005). The conservation of the APP sequence throughout phylogeny and its widespread expression suggest that it has important physiological roles. Several roles for APP were demonstrated in the brain. It was shown that it plays a role in the response of the cortex to loss of subcortical innervation (Wallace et al., 1993, 1995), neuroprotection (Mattson et al., 1993; Schubert and Behl, 1993), stimulation of neurite outgrowth (Small et al., 1994), and cellular adhesion (Breen et al., 1991), and it may function as a mitogenic growth factor (Saitoh et al., 1989) or neurotrophic factor (Araki et al., 1991; Milward et al., 1992; Qiu et al., 1995). In PC12 cells, APP potentiates the neurotrophic effects of nerve growth factor via activation of the IRS‐1 signaling pathway (Wallace et al., 1997). A recent study revealed a function for soluble APP as a regulator of subventricular zone (SVZ) progenitor proliferation in the adult CNS. The SVZ is the largest neurogenic area of the adult brain and it is also a major binding site for soluble APP (Caille et al., 2004). Furthermore, APP plays a role in controlling cell cycle progression during cortical development, particularly affecting G2 and mitosis (Lopez‐Sanchez et al., 2005). Proteolytic cleavage of the 695, 751, and 770 amino acids APP forms produces the 40–42 amino acids Ab (Glenner and Wong, 1984; Masters et al., 1985). This is the product of cleavage by two enzymes termed b‐ and g‐secretases. An alternative cleavage by a‐secretase precludes the production of Ab. Ab is the major component of the amyloid deposited in senile plaques and cerebral vessel walls of patients with AD, DS, familial and sporadic CAA and HCHWA‐D, and in nondemented aged individuals. Diseases that involve Ab deposition seem to be heterogeneous in etiology and clinical presentation. However, in some cases, abnormalities in APP proteolytic processing are a critical step in the molecular pathology of the disease (De Jonghe et al., 1998). Several APP missense mutations were identified within or flanking the Ab region in patients with HCHWA‐D and familial AD (FAD). These substitutions enhance the fibrillation of the peptide, increase the amount of the secreted peptide, or increase the proportion of secreted Ab ending at residue 42 and 43 rather than at 40 (Levy et al., 1990; Chartier‐Harlin et al., 1991; Goate et al., 1991; Murrell et al., 1991; Naruse et al., 1991; Yoshioka et al., 1991; Citron et al., 1992; Mullan et al., 1992a; Cai et al., 1993; Suzuki et al., 1994; Tamaoka et al., 1994; Younkin, 1994). Presenilin (PS1 and PS2) missense mutations found in some families with early onset FAD (Levy‐Lahad et al., 1995; Sherrington et al., 1995; Price and Sisodia, 1998) also modify the length of Ab generated (Borchelt et al., 1996; Duff et al., 1996; Citron et al., 1997). The longer Ab contains more hydrophobic residues than Ab1–40, and therefore it is more fibrillogenic and enhances aggregation of shorter peptides (Hilbich et al., 1991; Burdick et al., 1992; Jarrett and Lansbury, 1993; Jarrett et al., 1993). There are indications that Ab deposition causes neuronal death via a number of possible mechanisms including oxidative stress, excitotoxicity, energy depletion, inflammation, and apoptosis. Extensive neuronal loss and synaptic changes in the cerebral cortex and hippocampus as well as other areas of the brain essential for cognitive and memory functions are other hallmarks of AD. a1‐Antichymotrypsin. a1‐Antichymotrypsin (ACT) was detected in activated astrocytes during normal aging of humans and monkeys (Abraham et al., 1989) and in several neurodegenerative diseases (Abraham et al., 1990). It is present in both amorphous and classic plaques of AD, DS, and normally aged brains (Abraham and Potter, 1989; Abraham et al., 1991). ACT forms stable complexes with Ab1–42, comparable in

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specificity and stability to a protease/inhibitor interaction. Some studies have shown that ACT accelerates fibrillation of Ab1–42, increasing the neurotoxicity of the Ab peptide (Ma et al., 1996), while other studies report that it binds to Ab but inhibits Ab fibrillation (Fraser et al., 1993; Ma et al., 1994; Eriksson et al., 1995; Aksenova et al., 1996; Janciauskiene et al., 1996; Webster and Rogers, 1996; Hughes et al., 1998; Yamin et al., 1999; McLaurin et al., 2000). Similar controversial results have been reported for the effect of ACT on Ab neurotoxicity (Aksenov et al., 1996; Aksenova et al., 1996; Ma et al., 1996; Schubert, 1997; Mucke et al., 2000). Thus, ACT functions as a molecular chaperone to influence fibril formation through specific complexes with Ab. The increase of plaque load in transgenic mice overexpressing both APP and ACT compared to APP transgenic mice suggests that ACT promotes Ab deposition (Nilsson et al., 2004). Furthermore, murine apolipoprotein E (ApoE) and human ACT separately and synergistically facilitate both diffuse Ab‐immunoreactive deposition and fibrillar amyloid deposition, and thus also promote cognitive impairment in aged APP transgenic mice (Nilsson et al., 2004). It was recently shown that ACT also inhibits Ab degradation (Yamin et al., 1999; Abraham et al., 2000) by indirectly inhibiting a protease that degrades Ab (Yamin et al., 1999). The binding of Ab to ACT renders ACT inactive as a protease inhibitor. The loss of ACT inhibitory activity implies deregulation of protease(s) at foci of Ab biosynthesis, these proteases being elevated during inflammation associated with AD. Moreover, the complex of Ab1–42 with ACT structurally resembles that of serpin/protease complexes, and because the latter have biological activities beyond that of protease inhibition, it is possible that ACT/Ab1–42 complexes also have as yet undiscovered biological activities which contribute to self‐propagating neurotoxic pathologies. It is conceivable that ACT/protease complexes and/ or ACT/Ab complex may upregulate ACT biosynthesis, which by elevating local ACT levels in the presence of increasing levels of Ab could sustain pathological cycles (Janciauskiene et al., 1998). Blockage of ACT inhibitory activity due to its association with Ab could also result in upregulation of proteases that are targets of ACT. These could include proteases associated with Ab biosynthesis or degradation and proteases linked to cytokine activity. The ApoE gene is a major risk factor for developing AD and there are genetic data demonstrating that ACT polymorphism is likely to modify the role of the ApoE gene in AD. There are numerous genetic studies on the association of a polymorphism in the ACT gene with AD with varied results. A locus for familial early‐onset AD was identified on the long arm of chromosome 14, proximal to the aCt gene (Mullan et al., 1992b) and a polymorphism in its gene has been proposed to increase the risk of developing AD in association with the ApoE e 4/4 genotype (Wang et al., 2002). Furthermore, positive associations of ACT with CAA have been described (Yamada et al., 1998). However, this finding was not corroborated in a variety of familial and sporadic cases (Haines et al., 1996; Lamb et al., 1998). Investigation of the distribution of genotypes of ApoE and ACT in idiopathic Parkinson’s disease (PD) revealed that the ACT gene might be one of the susceptibility factors for PD without association with ApoE genotypes (Yamamoto et al., 1997; Wang et al., 2001; Lin et al., 2004). However, no association of the ACT gene and PD was found in other studies (Grasbon‐Frodl et al., 1999; Munoz et al., 1999; Tang et al., 2002). It was suggested that a consideration of differences in genetic background seems warranted when evaluating susceptibility factors for neurodegenerative diseases. Glia‐derived nexin (protease nexin I). Protease nexin I (PN‐1) is a neurite‐promoting factor synthesized and secreted by astroglial cells in embryos and may play a role in modeling and remodeling of brain tissues during development (Crisp et al., 2002). In adults, PN‐1 is found mainly in the brain (Monard, 1993) and it is believed that its primary role is a rapid thrombin inhibition and clearance during trauma and loss of vascular integrity (Crisp et al., 2002). The ability of astrocytes to renew the expression of PN‐1 after selective delayed neuronal death and the prolonged presence of the protease inhibitor in the immediate neighborhood of preserved neurons in brain areas where degeneration occurs indicate that it may play a role in structural rearrangements of the CNS (Fumagalli et al., 1999). Furthermore, PN‐1, ACT, as well as APP containing the Kunitz‐type domain participate in the structural stability of synaptic connections (Festoff et al., 2001). Transgenic mice with ectopic or increased expression of PN‐1 in postnatal neurons have altered synaptic transmission and plasticity (Luthi et al., 1997). These mice develop disturbances in motor behavior starting at 12 weeks of age, with some of the


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histopathological changes described in early stages of human motor neuron disease and neurogenic muscle atrophy in old age. In addition, sensorimotor integration is impaired. It is suggested that axonal dysfunction rather than cell death underlies these phenotypes (Meins et al., 2001). PN‐1 also appears to play a crucial role following ischemic injury (Hoffmann et al., 1992) and an upregulation of PN‐1 has been found in the peripheral nervous system following nerve lesion. It seems that tissue plasminogen activator (tPA) and its inhibitor, PN‐1, control the cell‐associated proteolytic activity in the injured nerve (Meier et al., 1989; Bleuel et al., 1995). Additional experiments suggest a neurotrophic role for PN‐1 (Oppenheim et al., 1993). Small doses of PN‐1 completely prevented axotomy‐induced spinal motoneuron death in neonatal mouse. A protease/inhibitor balance may be involved in regulating the fate of neuronal cells during development (Houenou et al., 1995). PN‐1 expression is regulated by neuronal activity and it plays a crucial role in the regulation of brain proteolytic activity and the functioning of sensory pathways. PN‐1 knockout mice present increased brain proteolytic activity, which is correlated with an activity‐dependent decrease in the NRI subunit of the N‐methyl‐D‐aspartate (NMDA) receptor, coupled to decreased sensory‐ evoked potentials in the barrel cortex and impaired whisker‐dependent sensory motor function (Kvajo et al., 2004). Immunohistochemical experiments showed that a small subset of neuritic plaques in brains affected by AD stain positively with an antibody to PN‐1 (Rosenblatt et al., 1989). It was also shown that the amount and activity of PN‐1 in brains of individuals with AD were reduced compared to control values suggesting that the decrease in PN‐1 is due to formation of PN‐1–protease complexes (Wagner et al., 1989). Strong immunoreactivity for PN‐1 was observed in capillaries and in smooth muscle cells of arteries and arterioles in the adult human cerebral cortex. Expression of PN‐1 was also abundant in astroglial processes in the parenchyma and in perivascular astroglial processes of the human cerebral cortex. PN‐1 around blood vessels might play a major protective role against extravasation of thrombin and possibly other serine protease into the human brain (Choi et al., 1990). The number of blood vessels exhibiting PN‐1 immunoreactivity, as well as PN‐1 activity, was markedly reduced in the brains of patients with AD compared to age‐matched controls. Thus, an imbalance between PN‐1 and thrombin may be a contributing factor in the pathology of AD (Vaughan et al., 1994; Shea, 1995). Neuroserpin. Neuroserpin is a member of the serpin family that is closely related in structure to another serpin, a1‐antitrypsin (Briand et al., 2001; Silverman et al., 2001). It is primarily expressed in neurons in the brain (Osterwalder et al., 1996) and is secreted from the axonal growth cones of the central and peripheral nervous system where it inhibits the enzyme tPA (Osterwalder et al., 1996; Hastings et al., 1997; Osterwalder et al., 1998; Barker‐Carlson et al., 2002; Belorgey et al., 2002). Neuroserpin expression pattern and its inhibitory activity suggest that it participates in neuronal plasticity and learning (Seeds et al., 1995), controlling axonal growth, regulating emotional behavior and memory, reducing epileptic seizure activity, and limiting damage in cerebral infarction (Yepes et al., 2000; Cinelli et al., 2001; Hill et al., 2002; Parmar et al., 2002; Yepes et al., 2002; Madani et al., 2003; Teesalu et al., 2004; Yepes and Lawrence, 2004). Mutations found in the neuroserpin gene result in polymerization and aggregation of the protein in neurotoxic inclusion bodies called Collins bodies (Davis et al., 1999a). Aggregation of the protein results in diminution in neuroserpin levels, with a potential increase in proteolysis and loss of neuronal function (Takao et al., 2000; Yazaki et al., 2001). Such mutations are associated with familial encephalopathy with neuroserpin inclusion bodies (FENIB) (Davis et al., 1999b). In this dementia, ordered polymers of neuroserpin are retained within the endoplasmic reticulum of neurons (Belorgey et al., 2002; Belorgey et al., 2004; Miranda et al., 2004). The number of inclusions is directly related to the rate of polymer formation and inversely proportional to the age of onset of the dementia. While the Syracuse mutant (Ser49Pro) causes dementia in middle age, the more rapidly polymerizing Portland mutant (Ser52Arg) causes an onset of dementia in the early twenties with the formation of higher amounts of inclusions (Belorgey et al., 2002; Davis et al., 2002; Belorgey et al., 2004; Miranda et al., 2004). Thus, a variant form of neuroserpin is yet another serpin mutant, which results in a conformational disease similar to angioedema resulting from polymerization of C1 inhibitor, thrombosis with polymerization of antithrombin, and emphysema with polymerization of ACT (Carrell and Lomas, 2002; Lomas and Mahadeva, 2002). Disorders characterized by polymerization

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of serpin mutants are termed serpinopathies. Other conformational dementias, with protein aggregation in the brain, involve Ab deposition in AD, accumulation of a‐synuclein (Lewy bodies) in PD, or of tau protein (Pick bodies) in some forms of familial frontotemporal dementia (Murrell et al., 1999; Tran and Miller, 1999; Walker and LeVine, 2000; Shimura et al., 2001). Plasminogen activator inhibitor. Plasminogen activator inhibitor (PAI) and its target proteases such as plasminogen activators and thrombin are involved in a variety of physiological and pathological processes in the brain. tPA is a serine protease, which is involved in neuronal plasticity and cell death induced by excitotoxins and ischemia in the brain. As a member of the serpin superfamily of serine protease inhibitors and an acute phase response component, PAI has potential roles in nervous system homeostasis and neuroprotection. Human astrocytes regulate capillary fibrinolysis in vitro by inhibiting tPA and enhancing type 1 PAI (PAI‐1) expression (Kim et al., 2003). PAI is regulated by the transforming growth factor‐b1 (TGF‐b1) in astrocytes. PAI‐1 produced by astrocytes mediates the neuroprotective effect of TGF‐b1 in NMDA‐induced neuronal cell death. Focal cerebral ischemia in mice induced a dramatic overexpression of PAI‐1, and the expression of PAI‐1, but not tPA, was increased 1 h after middle cerebral artery occlusion in the ischemic core (Docagne et al., 1999; Chang et al., 2003; Zhao et al., 2003). PAI‐1 in the CNS maintains the morphology of neurites via activation of the ERK‐related pathway in neurons (Soeda et al., 2004). An imbalance in the tPA/PAI‐1 system was found in disorders such as neurofilament 2 (NF2)‐associated schwannomas (Siren et al., 2004). Several lines of evidence implicate the tPA/plasmin system in AD. Accumulation of Ab depends on both its generation and clearance and the tPA/plasmin system is implicated in Ab degradation. Immunohistochemistry studies showed that tPA accumulates on senile plaques (Rebeck et al., 1995). In two different mouse models of AD, chronically elevated Ab peptide in the brain correlates with the upregulation of PAI‐1 and inhibition of the tPA/plasmin system. In addition, Ab injected into the hippocampus of mice lacking either tPA or plasminogen persists, inducing PAI‐1 expression and causing activation of microglial cells and neuronal damage. Conversely, Ab injected into wild‐type mice is rapidly cleared and does not cause neuronal degeneration (Melchor et al., 2003). These data suggest that the tPA/plasmin proteolytic system has a role in the clearance of Ab. Moreover, PAI inhibits Ab‐induced neurodegeneration (Melchor et al., 2003). Administration of PAI‐1 prevents the tPA‐ promoted neuronal degeneration in rat brain (Tsirka et al., 1996). DS patients, with trisomy 21, show overexpression of the Ab and tend to develop AD‐type pathology early in life (Masters et al., 1985; Wisniewski et al., 1985). Adults with DS have reduced levels of PAI in blood compared to control subjects. These reduced levels of PAI may explain the low incidence of atherosclerotic vascular disease in DS (Hopkins et al., 2000). Thus, the reduced activity of the tPA/plasmin proteolytic system has a role in the clearance of Ab, contributing to the progression of AD. Antithrombin. Thrombin is a key coagulation factor and its major inhibitors are antithrombin III (AT) in the plasma and PN‐1 in the brain. Thrombin has many other effects such as induction of brain edema, angiogenesis, and cell proliferation—three important factors for the prognosis of glioma. It may also play a role in ischemic brain damage. It was shown that it mediates hippocampal neuroprotection against ischemia at low concentrations but causes degeneration at high concentrations, thus determining neuronal cell death or survival after brain ischemia (Striggow et al., 2000; Hua et al., 2003a, b). Accordingly, antithrombin may be a potential neuroprotective agent as it was shown to have a positive effect on the recovery of incomplete spinal cord injury in rats (Arai et al., 2004; Hirose et al., 2004). Thrombin inhibition by argatroban attenuates neurodegeneration and cerebral edema formation following transient forebrain ischemia (Ohyama et al., 2001). Plasma thrombin/antithrombin III complex (TAT) levels are sensitive markers of the severity of experimental autoimmune encephalomyelitis (EAE), an animal model for human multiple sclerosis. TAT values increased immediately prior to the development of symptoms and decreased concurrently with the improvement of symptoms (Inaba et al., 2001). Pigment epithelium‐derived factor. Pigment epithelium‐derived factor (PEDF) is a member of the serpin superfamily, which has no inhibitory activity against any known proteases. Native bovine and recombinant human PEDF promote the survival and differentiation (neurite outgrowth) of embryonic chick spinal cord motor neurons in vitro in a dose‐dependent manner (Houenou et al., 1999).


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3.2 Cysteine Protease Inhibitors The cysteine proteases contain a sulfhydryl group in their active sites. This family encompass IL‐1b‐ converting enzyme (ICE) and a related protease, caspase (Black et al., 1989; Kostura et al., 1989), and calpain that plays a key role as a biomodulator in various cellular functions in response to Ca2þ‐mediated signaling (Saido et al., 1994; Sorimachi et al., 1994; Suzuki et al., 1995; Sorimachi et al., 1997; Ono et al., 1998). It also contains lysosomal cathepsins, synthesized as preproenzymes, which undergo activation by limited proteolysis. At least 11 human cathepsins are currently known (Turk et al., 2000) which have been divided in two families: Cathepsin L‐like (L, V, K, S, W, F, H) and cathepsin B‐like peptidases (Karrer et al., 1993; Berti and Storer, 1995). While most of the enzymes, including cathepsin L are endopeptidases, cathepsin B exhibits an exopeptidase activity (Musil et al., 1991). Their release into extracellular fluid can cause proteolytic tissue damage leading to multiple organ failure (Jochum et al., 1994). They also participate in the matrix destruction associated with inflammation, tumor invasion, and metastasis (Nomura and Katunuma, 2005). Endogenous inhibitors are important for regulation of mature enzymes. Those include the cystatins (Turk and Bode, 1991; Twining, 1994; Turk et al., 1997) and thyropins (Lenarcic and Bevec, 1998), as well as the nonspecific protease inhibitor a2‐M (Mason, 1989). On the basis of sequence similarity, the cystatin superfamily has been subdivided into three families: stefins, cystatins, and kininogens (Barrett et al., 1986). Three residues, Gly11, Gln55, and Gly59 (cystatin C numbering) are conserved in all the cystatins and have an important role in inhibition (Turk and Bode, 1991; Turk et al., 1997). Stefins are single‐chain proteins of about 100 amino acids that lack carbohydrate and disulfide bonds. They are primarily intracellular, found in various cells and tissues in animals, but are also secreted into extracellular fluids (Abrahamson and Grubb, 1994). Stefins A and B, also called cystatins A and B, do not contain secretory signal. They were found in mammals, including rats, mice, bovine, and humans (Turk et al., 1997). Stefin C was identified in bovine (Turk et al., 1993) and stefin D in pigs (Lenarcic et al., 1996). Members of the cystatin family are composed of about 120 amino acids. They are usually nonglycosylated single‐chain proteins having a secretory signal, two intramolecular disulfide bridges, and a binding region. They are widely distributed in nature including lower organisms (Rawlings and Barrett, 1990; Turk et al., 1997). The family consists of cystatin C, cystatin S, cystatin D (Freije et al., 1991), cystatin E/M (Ni et al., 1997; Sotiropoulou et al., 1997), cystatin F (Abrahamson and Grubb, 1994), and cystatin G (Abrahamson et al., 2003). Related to this family is the cystatin‐related epididymal spermatogenic (CRES) subfamily, containing cystatins expressed in the male reproductive tract which lack some of the consensus sites necessary for the cystatin inhibition of C1 cysteine proteases (Cornwall and Hsia, 2002). Several additional genes belonging to this subfamily have been identified, such as mouse Cymg1 (Xiang et al., 2004; Xiang et al., 2005), mouse cystatin E1 and E2 (Li et al., 2003), mouse cystatin SC and TE‐1 (Li et al., 2002), Macaca mulatta cystatin gene 11 (CST11) (Hamil et al., 2002), the mouse cystatin M/E ortholog (Zeeuwen et al., 2002), and human cystatin‐like molecule (CLM) (Sun et al., 2003). Cystatin E/M differs from other cystatins in biochemical properties, chromosomal localization, and by having a more tissue‐specific distribution. Unlike the other cystatins, it exists as a glycosylated protein in addition to the nonglycosylated form (Ni et al., 1997). It is expressed in the brain, neuronal cells, and lungs, and may play a role in neuronal development (Hong et al., 2002). A null mutation of the cystatin M/E gene causes juvenile lethality and defects in epidermal cornification (Zeeuwen et al., 2002). Most of the cystatin family genes, including the cystatin C gene, and six to eight genes for cystatin S are clustered on the short arm of chromosome 20 (Thiesse et al., 1994), whereas the cystatin E/M gene is located on chromosome 13 (Stenman et al., 1997). CLM is ubiquitously expressed and may have a role in hematopoietic differentiation or inflammation (Sun et al., 2003). Cystatin F is expressed almost exclusively in immune cells and it could have a role regulating papain‐like cathepsins involved in antigen presentation or in cancer progression (Langerholc et al., 2005). Salivary cystatins may be involved in immune response through the cytokine network (Kato et al., 2004). Cystatin D is a secreted inhibitor found in human saliva and tears. It has a restricted inhibition profile (Alvarez‐Fernandez et al., 2005). Cystatin M is a novel candidate tumor suppressor gene for breast cancer (Shridhar et al., 2004; Zhang

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et al., 2004). Recently, fetuin B gene was identified as a member of a cysteine‐like gene family in mouse. Overexpression of fetuin B in skin squamous carcinoma cells led to suppression of tumor growth in nude mice (Hsu et al., 2004). Mouse cystatin 10 is expressed in cartilage and it is implicated in endochondral ossification (Koshizuka et al., 2003). Cystatin C (Bobek and Levine, 1992), also known as g trace (Hochwald et al., 1967), is found in all mammalian body fluids and tissues (Bobek and Levine, 1992). In vitro experiments have indicated that it can inhibit cathepsins B, H, L, and S (for review (Bernstein et al., 1996)). In addition to being a protease inhibitor, cystatin C itself is a target of proteolysis (Rudensky et al., 1991; Rider et al., 1996) and is inactivated by proteolytic degradation by cathepsin D and elastase (Abrahamson et al., 1991; Lenarcic et al., 1991). It has a broad spectrum of biological roles ranging from antiviral to antibacterial properties (Bobek and Levine, 1992), bone resorption, (Lerner and Grubb, 1992), tumor metastasis (Huh et al., 1999; Taupin et al., 2000), modulation of inflammatory responses (Warfel et al., 1987; Bobek and Levine, 1992), and cell proliferation and growth (Sun, 1989; Tavera et al., 1992). Similar to a variety of activities that have been associated with other protease inhibitors in the brain (Sun, 1989; Akopyan, 1991; Lee et al., 1991; Hoffmann et al., 1992; Roberts‐Lewis et al., 1993), cystatin C has been implicated in the processes of neuronal degeneration and repair of the nervous system. Cystatin C is also involved in astrocytic differentiation during mouse brain development (Kumada et al., 2004). Cystatin B (also called stefin B) is a cytosolic inhibitor of cysteine proteases including cathepsins B, H, L, and S (Barrett, 1986; Barrett et al., 1986; Turk and Bode, 1991). Data suggest that it is involved in neuroprotection by inhibiting cytosolic hydrolases. Furthermore, cystatin B is present in embryonic and adult neural stem cells and in the neuroepithelium, expressed by both neurons and glial cells differentiated from neural stem cells and in hippocampal cultures. It localized mainly to the nucleus in neural stem cells and in neurons, while in astrocytes cystatin B was also localized in the cytoplasm and in the lysosomes in glial cells. The presence of cystatin B in the nucleus of neural stem cells and in neurons suggests a novel function for this molecule (Brannvall et al., 2003). Kininogens are large multifunctional glycoproteins circulating in mammalian blood, and were initially identified as parent molecules for the vasoactive peptides, the kinins. They contain three copies of the cystatin family sequences and have up to nine disulfide bonds. They are divided into three groups: high‐ molecular weight kininogen (HK), low‐molecular weight kininogen (LK), and T‐kininogen (TK), the latter found only in rats (DeLa Cadena and Colman, 1991). Both human HK and LK are products of the same gene by alternative mRNA splicing (Kitamura et al., 1985). This gene is located on the long arm of chromosome 3 (Cheung et al., 1992). Kininogens are precursor molecules for the vasodilator kinin peptides: bradykinin and kallidin. Kallikreins are the specific activators of kininogens. The activation of the kinin system is particularly important in blood pressure regulation and in inflammatory reactions (Agostoni and Cugno, 2001). In addition to being a source of bradykinin, HK also functions as a factor in the blood coagulation cascade (DeLa Cadena and Colman, 1991). Complete absence of kininogens is characterized by prolonged clotting time (Stormorken et al., 1990). The kinin system is involved in many clinical situations including respiratory allergic reactions, septic shock, hypertension and its treatment, hypotensive transfusion reactions, heart diseases, pancreatitis, hereditary and acquired angioedema, AD, and liver cirrhosis with ascites (Agostoni and Cugno, 2001). It is also related to dermal diseases (Schremmer‐Danninger et al., 2004), spermatogenesis (Blaukat, 2003), diabetes (Damas et al., 2004), tumor angiogenesis, and tumor growth (Ikeda et al., 2004). Uncontrolled proteolysis as a result of imbalance between active proteases and their endogenous inhibitors has been associated with different diseases such as AD, cancer, rheumatoid arthritis and osteoarthritis, multiple sclerosis, muscular dystrophy, pancreatitis, liver disorders, lung disorders, lysosomal disorders, inflammation, Batten’s disease, diabetes, pycnodysostosis, periodontitis, myocardial disorders, and many others (Sohar et al., 1988; Assfalg‐Machleidt et al., 1990; Buttle et al., 1991; Delaisse et al., 1991; Nakamura et al., 1991; Trabandt et al., 1991; Lah et al., 1993; Bever and Garver, 1995; Calkins and Sloane, 1995; Kabanda et al., 1995; Thomssen et al., 1995; Duffy, 1996). There is considerable evidence connecting lysosomal cysteine proteases with aging. It was shown that lysosomes in the aged rat brain are less stable, with significant leakage of lysosomal enzymes either as mature enzymes or as inactive precursors (Nakamura et al., 1989; Bi et al., 2000). Furthermore, in many diseases precursor forms of lysosomal enzymes have


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leaked from the lysosomes. These precursor forms are more stable than mature enzymes and not susceptible to inhibition by endogenous inhibitors (Kirschke et al., 1995; Kos et al., 2000; Turk et al., 2000). Cystatins are associated with several pathologies of the CNS including AD and neuronal degeneration, and recently cystatin C was reported as a potential CSF marker for the diagnosis of Creutzfeldt–Jakob disease (Sanchez et al., 2004).

3.2.1 Cystatin C in the Nervous System and Neurological Disorders Hereditary Cerebral Hemorrhage with Amyloidosis, Icelandic Type Hereditary cerebral hemorrhage with amyloidosis, Icelandic type (HCHWA‐I) (Arnason, 1935; Gudmundsson et al., 1972), also called hereditary cystatin C amyloid angiopathy (HCCAA) (Olafsson et al., 1996), is an autosomal dominant form of CAA. Amyloid deposition in cerebral and spinal arteries and arterioles leads to recurrent hemorrhagic strokes causing serious brain damage and eventually fatal stroke before the age of 40 (Gudmundsson et al., 1972). While amyloid is also present in other tissues, such as skin and lymph nodes (Benedikz et al., 1990), the deposition in the CNS vasculature is particularly pronounced. The amyloid deposited in brain vessel walls is composed mainly of a variant of cystatin C. Comparison of the genes encoding cystatin C, isolated from normal tissue and from the brain of an HCHWA‐I patient, revealed a mutation in the Icelandic gene (Levy et al., 1989) that segregates with the disease in every case (Palsdottir et al., 1988; Abrahamson et al., 1990). The amyloid protein isolated from leptomeninges of HCHWA‐I patients starts at position 11 of normal cystatin C and has an amino acid substitution, Leu to Gln, at position 68 (L68Q) (Cohen et al., 1983; Ghiso et al., 1986). The mutation found in the cystatin C gene in HCHWA‐I patients was identified also in the cystatin C gene of a Croatian man with CAA and intracerebral hemorrhage (Graffagnino et al., 1995). The Leu68Gln variant cystatin C is less stable than the wild‐type protein and more prone to dimerization (Abrahamson and Grubb, 1994; Ekiel and Abrahamson, 1996; Wei et al., 1998). Prevention of domain swapping inhibits dimerization and amyloid fibril formation of cystatin C (Nilsson et al., 2004). Alzheimer’s Disease and Aging There are several indications that cystatin C may have a role in AD. Genetic data were presented demonstrating linkage of the cystatin C gene, localized on chromosome 20 (Abrahamson et al., 1989; Saitoh et al., 1989), with B/B homozygosity associated with an increased risk of developing late‐onset AD (Crawford et al., 2000; Finckh et al., 2000). While some studies were unable to replicate these findings (Maruyama et al., 2001; Roks et al., 2001; Dodel et al., 2002), the linkage was supported by others (Beyer et al., 2001; Olson et al., 2002; Goddard et al., 2004). The polymorphism in the cystatin C gene results in an amino acid exchange, which alters the hydrophobicity profile of the signal sequence (Finckh et al., 2000), resulting in a less efficient cleavage of the signal peptide and thus a reduced secretion of cystatin C (Benussi et al., 2003). Immunohistochemical studies revealed colocalization of cystatin C with Ab predominantly in amyloid‐ laden vascular walls and in senile plaque cores of amyloid in brains of patients with AD, DS, HCHWA‐D, and cerebral infarction (Maruyama et al., 1990; Vinters et al., 1990; Itoh et al., 1993; Haan et al., 1994; Levy et al., 2001). Cystatin C also colocalizes with Ab amyloid deposits in the brain of nondemented aged individuals (Levy et al., 2001), aged rhesus and squirrel monkeys (Wei et al., 1996), and transgenic mice overexpressing human bAPP (Levy et al., 2001; Steinhoff et al., 2001). Furthermore, immunohistochemical studies have shown dual staining with antibodies to Ab and to cystatin C in the same subpopulation of pyramidal neurons suggesting that Ab accumulates in a specific population of neurons, the same cell type in which cystatin C is highly expressed (Levy et al., 2001). Analysis of the association of cystatin C and Ab demonstrated a specific, saturable, and high‐affinity binding between cystatin C and both Ab1–42 and Ab1–40. Notably, cystatin C association with Ab results in a concentration‐dependent inhibition of Ab amyloid fibril formation, suggesting a protective role for cystatin C in AD (Sastre et al., 2004). Morphometric analyses of brains from patients with AD have shown that pyramidal cells in the prefrontal cortex and hippocampus displayed the highest activation of hydrolase‐positive vacuolar compartments (Cataldo et al., 1996). The neuronal staining of cystatin C in brains from patients with AD was primarily limited to pyramidal neurons in cortical layers duplicating the pattern of neuronal susceptibility

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in AD brains: the strongest staining was found in the entorhinal cortex, in the hippocampus, and in the temporal cortex; fewer pyramidal neurons were stained in the frontal, parietal, and occipital lobes (Deng et al., 2001). Immunostaining of cystatin C within neurons showed a punctate distribution, which colocalized with the endosomal/lysosomal protease cathepsin B (Deng et al., 2001). Upregulation of cathepsin synthesis in AD neurons and accumulation of hydrolase‐laden lysosomes indicate an early activation of the endosomal/lysosomal system in vulnerable neuronal populations, possibly reflecting early regenerative or repair processes (Cataldo et al., 1995, 1996). These neuropathological observations reinforce an association between cystatin C and cathepsin B and cathepsin D in AD and support a model of cystatin C involvement in the process of neuronal death in AD. Neuronal Degeneration Cystatin C has been implicated in the processes of neuronal degeneration and repair of the nervous system. Enhanced cystatin C expression was observed in response to injury, including facial nerve axotomy (Miyake et al., 1996), perforant path transections (Ying et al., 2002), hypophysectomy (Katakai et al., 1997), transient forebrain ischemia (Palm et al., 1995; Ishimaru et al., 1996), and induction of epilepsy (Aghajanyan et al., 1988; Peppard and Knap, 1999; Aronica et al., 2001; Hendriksen et al., 2001; Lukasiuk et al., 2002). Oxidative stress also stimulates an increase in cystatin C expression in cultured neurons (Nishio et al., 2000) and in cerebral microvascular smooth muscle cells (Wang et al., 2002), suggesting a role in regulation of apoptosis. Recently it was reported that cystatin C prevents degeneration of rat nigral dopaminergic neurons, involved in PD pathology (Xu et al., 2005). Cystatin C dimers bind to neuroglobin, a globin that protects neurons from hypoxia (Wakasugi et al., 2004). Moreover, a glycosylated form of rat cystatin C was found to be an autocrine/paracrine factor, required for the mitogenic activity of basic fibroblast growth factor on neural stem cells (Taupin et al., 2000). Several hypotheses can be envisioned to explain the involvement of cystatin C in the brain: it can stimulate cell proliferation, promote mitogenic activity of the cell, promote survival, or prevent cell death. These functions may involve inhibition of cathepsins. Lysosomal cathepsins are also involved in neuronal death either as initiators or direct agents (Cataldo and Nixon, 1990). Intense cytoplasm labeling of cathepsin B was detected when neurons had become morphologically altered with obvious shrinkage of the cytoplasm (Hill et al., 1997). Enhanced expression of several cathepsins has been documented in response to injuries similar to those inducing cystatin C expression upregulation, such as in transient ischemia (Nitatori et al., 1995; Yamashima et al., 1998). Furthermore, inhibitors of cathepsins B and L reduce neuronal damage in the hippocampus after ischemia (Tsuchiya et al., 1999). Although cathepsins B and D are typically localized in lysosomes, they can be released through exocytosis, and activated microglia secrete several proteases including cathepsin B (Buck et al., 1992), which can trigger neuronal apoptosis (Bannerman et al., 1998; Levkau et al., 1998; Kingham and Pocock, 2001). Some evidence suggests that, in degenerating neurons, cathepsins can be released into the cytoplasm and neuropil after disruption of lysosomes (Cataldo and Nixon, 1990; Cataldo et al., 1990; Cataldo et al., 1994; Roberg and Ollinger, 1998; Kagedal et al., 2001; Bidere et al., 2003; Boland and Campbell, 2004). Cystatin C is secreted from microglia (Zucker‐Franklin et al., 1987) and is significantly downregulated following microglial and macrophage activation (Warfel et al., 1987; Chapman et al., 1990). These data suggest that intravesicular, intracellular, or extracellular cathepsins and their inhibitor cystatin C are involved in neurodegeneration, and imbalance in their expression may cause or exacerbate the neuropathology. Brain Tumors Cathepsins B, H, and L have been shown to participate in processes of tumor growth, vascularization, invasion, and metastasis. These processes depend on replication of cells and their ability to adhere and degrade extracellular matrix, enabling migration. In vitro studies demonstrated that secreted cysteine proteinases are able to degrade basement membrane components (Boike et al., 1992). Active forms of cathepsin B and L, normally restricted to the lysosomal compartments inside mammalian cells, are found in the extracellular space in tumors, either in a soluble state (Poole et al., 1978) or bound to the plasma membrane (Sloane et al., 1986). Enzymatically active extracellular cathepsin B may be produced in tumors by self‐activation of the latent precursor of the enzyme (Mach et al., 1994). The levels of extracellular


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cathepsins and their endogenous inhibitors stefins and cystatin C can provide useful clinical information to predict disease‐free and overall survival in breast, lung, colorectal, brain, and head and neck cancer patients (Kos et al., 2000). Cathepsin B was the first cysteine protease identified in relation to malignant progression of human gliomas (McCormick, 1993; Sivaparvathi et al., 1995). Overexpression of cathepsin B was found in glioma cell lines and tumors (Rempel et al., 1994; Sivaparvathi et al., 1995), and its level in human glioma is associated with poor clinical symptoms (Kos and Lah, 1998; Strojnik et al., 1999). Cathepsins D, L, and H are also upregulated in the malignant progression of gliomas (Sivaparvathi et al., 1996 b, c, d). Conversely, cysteine protease inhibitors activities were reduced in human glioma and meningioma. In vitro studies have shown that inhibition of cysteine proteases reduces glioma invasion (Demchik et al., 1999). Progressive reductions in levels of cystatin C with corresponding increases in the malignancy of glioma cell lines were observed (Konduri et al., 2002). Furthermore, decreases in the activities of cysteine protease inhibitors may contribute to the malignant properties of brain tumors (Sivaparvathi et al., 1996a). These results implicate cystatin C in the invasiveness of human glioblastoma cells and suggest that sense transcripts of cystatin C may prove useful in cancer therapy (Konduri et al., 2002).

3.2.2 Cystatin B in the Nervous System and Neurological Disorders Progressive Myoclonus Epilepsies Progressive myoclonus epilepsies (PME) are inherited diseases characterized by myoclonic seizures, generalized epilepsy, and progressive neurological deterioration, particularly dementia and ataxia. Unverricht–Lundborg disease (EPM1) is one of the five major types of PME. It is a rare autosomal recessive disease caused by loss‐of‐function mutations in the gene encoding cystatin B (Pennacchio et al., 1996; Lafreniere et al., 1997; Lalioti et al., 1997). In most cases the gene contains large expansions of a dodecamer sequence (CCC CGC CCC GCG) located upstream of the 50 transcription start site of the cystatin B gene (Lalioti et al., 1997). While normal controls have 2–3 repeats of this sequence, patients have 30–75 repeats (Lalioti et al., 1998; Lalioti et al., 2003), resulting in markedly reduced cystatin B mRNA (Lalioti et al., 1998; Lalioti et al., 1999) and activity in lymphoblastoid cells (Rinne et al., 2002). The altering of the spacing of transcription factor‐binding sites from each other and/or the transcription initiation site due to repeat expansion is among the causes of reduction in cystatin B expression, and thus occurrence of EPM1 (Alakurtti et al., 2000). Cystatin B‐deficient mice develop myoclonic seizures and ataxia, similar to symptoms seen in the human disease (Pennacchio et al., 1998). Degenerating neurons were found in cerebellar granule cells (Pennacchio et al., 1998) and within the hippocampal formation and entorhinal cortex of mutant mice (Shannon et al., 2002). The mice also showed gliosis and increased expression of apoptosis and glial activation genes (Lehesjoki, 2003). Neuronal atrophy seems to be an important consequence of cystatin B deficiency independent of seizure events, suggesting a physiological role of this protein in maintenance of normal neuronal structure and survival (Shannon et al., 2002). In order to identify the cathepsins that contribute to EPM1, three candidate cathepsins were removed from cystatin B‐deficient mice and tested for rescue of their EPM1 phenotypes. Whereas removal of cathepsins L or S did not ameliorate any aspect of the EPM1 phenotype, removal of cathepsin B resulted in a reduction of cerebellar granule cell apoptosis depending on the age of the mouse. The incidence of an incompletely penetrant eye phenotype was also reduced upon removal of cathepsin B. The apoptosis and eye phenotypes were not abolished completely and the ataxia and seizure phenotypes experienced by cystatin B‐deficient animals were not diminished suggesting that another protein besides cathepsin B is also responsible for the pathogenesis, or that another protease can partially compensate for cathepsin B function (Houseweart et al., 2003). Brain Tumor It has been reported that cystatin B is highly expressed in glioblastoma multiforme tissues

compared to nonmalignant brain. It may be used as a diagnostic or functional marker for glioblastoma tumors (Zhang et al., 2003).

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3.2.3 Cystatin A in the Nervous System and Neurological Disorders Alzheimer’s Disease Immunohistochemical studies of human postmortem brain showed that cystatin A is expressed in a large population of neurons and a few cells surrounding cerebral blood vessels (pericytes). It was found in many neuritic plaques in AD patients (Bernstein et al., 1994), suggesting a role in amyloid deposition.

3.3 Metalloprotease Inhibitors Metalloproteases are endopeptidases that contain zinc. They are type II integral membrane proteins, including neutral endopeptidase (NEP); the endothelin converting enzymes (ECE), which catalyze the final step in the biosynthesis of the vasoconstrictor peptide; endothelin; the PEX gene product, associated with X‐linked hypophosphatemic rickets (Turner and Tanzawa, 1997); and matrix metalloproteases (MMPs). The MMP family is mainly composed of collagenases, gelatinases, and stromelysins. They are involved in degradation of extracellular matrix components (Matrisian, 1992; Birkedal‐ Hansen et al., 1993; Nagase, 1997) during normal connective tissue development as well as in pathologic processes such as destruction of joint tissue (Woessner, 1991; Cawston, 1995), tumor invasion, and metastasis (Folgueras et al., 2004; Mook et al., 2004; Ramnath and Creaven, 2004; Zucker and Vacirca, 2004). The metalloprotease inhibitors are called tissue inhibitors of metalloproteases (TIMPs). They share common structural features, including the characteristic six‐loop, resulting from 12 conserved cysteine residues forming intrachain disulphide bonds. The network of disulfide bonds provides compactness and rigidity to the molecule (Woessner, 1991). However, the four TIMPs (TIMP‐1, ‐2, ‐3, ‐4) have distinct properties and functions (Chambers and Matrisian, 1997). The genes encoding them have distinct chromosomal locations, producing proteins of 21–28 kDa. While TIMP‐1, ‐2, and ‐4 are freely diffusible, TIMP‐3 is associated with the extracellular matrix (Cawston, 1995). TIMPs are found in all connective tissues in the body (Denhardt et al., 1993; Cawston, 1995), regulating destruction of extracellular matrix with differences in tissue distribution, and TIMP‐2, ‐3, and ‐4 are also expressed in the brain. TIMP‐2 and ‐4 are constitutively expressed and their promoter region lacks the AP1 sites that confer inducibility on TIMP‐1 and ‐3 (De Clerck et al., 1994; Hammani et al., 1996). It was demonstrated both in vitro and in vivo that TIMPs block the deleterious effects of elevated production and activation of MMPs (Thorgeirsson et al., 1982; Nakajima et al., 1987; Mignatti and Rifkin, 1993). MMPs and TIMPs are often secreted in similar quantities by the same cells and conditions that lead to increased expression of tissue MMPs also lead to upregulation of TIMPs. However, the different TIMPs are under individual control and they can be inactivated by a variety of proteases such as neutrophil elastase and trypsin (Itoh and Nagase, 1995). The balance between MMPs and TIMPs affect a broad range of cellular functions such as growth and proliferation, apoptosis, and angiogenesis (Chambers and Matrisian, 1997; Vu et al., 1998). Imbalance between these proteases and their inhibitors seems to be an important factor in the pathogenesis of tumor invasion and arthritis (Alvarez et al., 1990; Hill et al., 1993; Ellis et al., 1994).

3.3.1 TIMPs in the Nervous System and Neurological Disorders In recent years, data have accumulated indicating that the MMP/TIMP system is expressed in the nervous system, where it regulates neuroimmune interactions and plays a major role in pathophysiological processes. There are in vivo and in vitro studies that highlight the contribution of the MMP/TIMP system to developmental and physiological processes including cell migration, axonal sprouting, and neuronal plasticity and to various diseases of the nervous system, involving blood brain–barrier breakdown, neuroinflammation, glial reactivity, neuronal death, and reactive plasticity (Giraudon et al., 2003; Khrestchatisky et al., 2003).


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3.3.2 Brain Tumors Multiple studies have demonstrated that MMPs are involved in the pathophysiology of gliomas. MMPs are a requirement for glioma growth and overexpression or deregulation of some MMPs is associated with a poorer clinical outcome (Rao et al., 1993, 1996; Nakagawa et al., 1994; Forsyth et al., 1998, 1999). It was shown that a synthetic inhibitor of MMP inhibits glioma proliferation, tumor growth, and vascularity in an in vivo model of glioma (Price et al., 1999). MMPs‐1, ‐2, ‐3, ‐7, and ‐9 (Apodaca et al., 1990; Nakano et al., 1993; Rao et al., 1993, 1996; Costello et al., 1994; Nakagawa et al., 1994, 1996; Nakano et al., 1995; Sawaya et al., 1996; Yamamoto et al., 1996; Forsyth et al., 1998, 1999; Price et al., 1999; Raithatha et al., 2000), and their inhibitors TIMPs 1–4 (Nakagawa et al., 1994; Matsuzawa et al., 1996; Lampert et al., 1998; Mohanam et al., 1998) are found in glioma cell lines and gliomas and in tumor vasculature observed in surgical specimens. MMPs‐2 and ‐9 are involved in invasion and angiogenesis. Overexpression of TIMP‐1 in a glioma cell line resulted in reduced invasion in vitro but also decreased proliferation (Matsuzawa et al., 1996). Furthermore, it was shown that their expression and activity are higher for all gliomas, irrespective of grade, than in normal brains (Forsyth et al., 1998, 1999). While it was demonstrated that TIMPs could inhibit invasion and sometimes metastases (Hicks et al., 1984; Schultz et al., 1988; Albini et al., 1991; DeClerck et al., 1991; Ponton et al., 1991; Khokha et al., 1992a, b; Montgomery et al., 1994), inconsistencies exist regarding the levels of TIMPs in gliomas. Some investigators found a reduced expression of TIMP‐1 and ‐2 (Mohanam et al., 1995) with increasing glioma grade, while others (Nakagawa et al., 1994; Lampert et al., 1998) found an upregulation of TIMPs‐1 and ‐2. However, it is hypothesized that an imbalance between MMPs and their inhibitors resulting from a loss of specific TIMP expression allows the malignant progression of gliomas.

3.3.3 Alzheimer’s Disease There are several studies suggesting that MMPs and their inhibitors are involved in the pathogenesis of AD. Studies of brain samples of AD and elderly patients used as controls for TIMP immunoreactivity have shown that TIMP staining was localized to neuritic senile plaques, neurofibrillary tangles, and Purkinje cells (Peress and Perillo, 1995). It was demonstrated that an integral membrane MMP cleaves APP at the a‐secretase domain (Roberts et al., 1994; Hooper et al., 2000). More specifically, APP contains a protease inhibitor domain for the matrix metalloproteinase gelatinase A that has an APP secretase‐like activity, with a‐secretase characteristics (Miyazaki et al., 1993). Several MMPs and their inhibitors are also implicated in the degradation of Ab. MMPs such as neprilysin (Iwata et al., 2000; Reilly, 2001; Yasojima et al., 2001; Apelt et al., 2003; Clarimon et al., 2003; Helisalmi et al., 2004; Sakai et al., 2004; Yamada, 2004), insulin‐degrading enzyme (Perez et al., 2000; Leissring et al., 2003), and ECE‐1 (Eckman et al., 2001) were found to be Ab‐degrading enzymes prone to inhibition by MMP inhibitors. It was shown that MMP (EC 3.4.24.15) (MP24.15) activates Ab‐degrading serine proteases. Because the serpin inhibitor ACT blocked Ab degradation by MP24.15 it was suggested that ACT may cause Ab accumulation by inhibiting an Ab‐degrading enzyme or by direct binding to Ab, rendering it degradation resistant (Yamin et al., 1999). These observations suggest that downregulation of MMPs activity is likely to be related to AD pathology and to the Ab accumulation associated with normal aging.

3.3.4 Vascular Dementia Increased expression of brain MMPs and TIMPs were found in cerebrovascular disease. Patients with multiinfarct and small‐vessel vascular dementia (VaD) have elevated levels of MMP‐9 in the CSF compared with AD patients and controls. Although increased MMP‐9 in CSF is not specific for VaD, it could provide an additional biological marker for the diagnostic discrimination of patients with VaD and AD (Adair et al., 2004).

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3.3.5 Multiple Sclerosis Abnormality in the inhibition of MMPs may play an etiological role also in multiple sclerosis. Average serum gelatinase B, TIMP‐1, and TIMP‐2 levels were significantly higher in multiple sclerosis patients than in healthy controls. While gelatinase B levels were significantly higher during clinical relapse compared with periods of clinical stability, there was a trend for TIMP‐2 levels to be lower during relapse compared with nonrelapse periods (Lee et al., 1999).

3.4 Aspartic Protease Inhibitors Mammalian aspartic proteases are endogenous, containing aspartic residues in their active sites (McDonald, 1985). Most of them belong to the pepsin family, including gastric proteases, such as pepsins and chymosins, and nongastric proteases such as rennin (processing enzyme), lysosomal cathepsin D, and nonlysosomal cathepsin E (Neurath, 1989). The inhibitors of aspartic proteases are the pepstatins, first isolated in the early 1970s by Umezawa et al (1970) from culture fluid of Streptomyces strains. Pepstatins are pentapeptides containing a unique amino acid sequence, namely, AHMHA or statin, which is important for the inhibitory activity (Umezawa, 1982). The most potent and specific reversible inhibitors of cathepsin D are pepstatins containing an isovaleryl group (Umezawa, 1982), including pepstatin A. Pepstatin A also inhibits other carboxyl proteases, such as pepsin and rennin, but does not inhibit trypsin, chymotrypsin, papain, and plasmin (Umezawa, 1982). It was shown to play a role in a variety of disorders, including muscular dystrophy (Schorr et al., 1978), myocardial ischemia (Wildenthal, 1978; Kalra et al., 1989), microbial infections (Tsuobi et al., 1985; Borg and Ruchel, 1988), HIV (Seelmeier et al., 1988; Grinde et al., 1989), and cancer invasion and metastasis (Giraldi et al., 1977; Briozzo et al., 1988; Leto et al., 1990; Rochefort et al., 1990; Tandon et al., 1990).

3.4.1 Pepstatin in the Nervous System and Neurological Disorders g‐Secretase activity is inhibited by pepstatin and more potently, by an aspartyl protease transition‐state analog inhibitor that blocks formation of Ab in mammalian cells. Several studies suggest that g‐secretase activity is catalyzed by a PS1‐containing macromolecular complex (Li et al., 2000; Xia et al., 2000; Zhang et al., 2001). In vitro studies have shown binding of pepstatin to both PS1 and PS2 (Evin et al., 2001). PS1 and PS2 missense mutations (Levy‐Lahad et al., 1995; Sherrington et al., 1995; Price and Sisodia, 1998) cause early onset FAD by altering g‐secretase cleavage, elevating levels of the more amyloidogenic Ab42(43) but not Ab40 (Borchelt et al., 1996; Duff et al., 1996; Citron et al., 1997). PS1 transgenic mouse models do not develop Ab deposition upon aging. However, when crossed to APP overexpressing mice, PS1 mutants accelerate the accumulation of amyloid and show a considerably enhanced amyloid phenotype compared to the parental APP mice, most likely due to the influence of elevated Ab42(43) on deposition rates (Borchelt et al., 1997; Holcomb et al., 1998). An aspartyl protease and its inhibitor are also involved in the degradation of Ab as was demonstrated using postmortem human and fresh rat brains. The highest Ab degrading activity in the soluble fractions occurred between pH 4–5, and this activity was inhibited by pepstatin, implicating an aspartyl protease (McDermott and Gibson, 1997).

Alzheimer’s Disease

This chapter presented data exemplifying the significance of controlled balance between proteases and their inhibitors in the brain. Factors that affect the levels of certain proteases or their inhibitors result in imbalance between the two, giving rise to multiple pathologies observed in aging and various diseases.

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Neuropeptidases

K.‐S. Hui

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 626

2 2.1 2.2 2.3 2.4 2.5 2.6

Abnormalities of Neuropeptides and Neuropeptidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 626 Neuropeptides in Degenerative Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 626 Neuropeptides in Cerebrospinal Fluid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 627 Neuropeptides in Neurological Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 628 Neuropeptide Degradation in Neurological Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 628 Exopeptidases in Aging and in Neurological Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 629 Endopeptidases in Neurological Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 629

3

Classification of Exo‐ and Endopeptidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 630

4 4.1 4.2 4.3 4.4

Enkephalin‐Processing Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 630 Prohormone Thiol Protease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 632 Prohormone Convertase 1 and 2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 632 Carboxypeptidase H/E . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 632 Arg/Lys Aminopeptidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 633

5 5.1 5.2 5.3 5.3.1 5.3.2 5.3.3

Enkephalin‐Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 633 Endopeptidase 24.11 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 634 Angiotensin‐Converting Enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 634 Aminopeptidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Aminopeptidase N . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Puromycin‐Sensitive Aminopeptidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Neuron‐Specific Aminopeptidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 638

6 6.1 6.2 6.3

Other Peptidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641 Tripeptidyl Peptidase (TPP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641 N‐Acetylated a‐Linked Acidic Dipeptidase (NAALADase) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641 Bestatin‐Insensitive Aminopeptidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 642

7 7.1 7.2 7.3 7.4 7.5

Therapeutic Uses of Neuropeptidase Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 642 Synthetic Anti‐Enkephalinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 642 Synthetic Anti‐Aminopeptidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 642 Endogenous Peptidase Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 643 Endogenous Anti‐Aminopeptidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 643 Inactivation of Arg0‐Met‐Enk . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 643

8

Neuropeptidases Regulated by Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 644

9

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 644

#


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Abstract: Neuropeptides are neurotransmitters and modulators distributed in the central nervous system (CNS) and peripheral nervous system. Their abnormalities cause neurological and mental diseases. Neuropeptidases are enzymes crucial for the biosynthesis and biodegradation of neuropeptides. We here focus on the peptidases involved in the metabolism of the well‐studied opioid peptides. Bioactive enkephalins are formed from propeptides by processing enzymes—prohormone thiol protease, prohormone convertase 1 and 2 (PC 1 and 2), carboxypeptidase H/E, and Arg/Lys aminopeptidase. After they exert their biological effects, enkephalins are likely to be inactivated by degrading enzymes—angiotensin‐converting enzyme (ACE), aminopeptidase N (APN), puromycin‐sensitive aminopeptidase (PSA), and endopeptidase 24.11. Recently, a neuron‐specific aminopeptidase (NAP), which was a putative enkephalin‐inactivating enzyme at the synapses, was found. Neuropeptidases are useful drug targets and their inhibitors can be therapeutic. Synthetic anti‐enkephalinases and anti‐aminopeptidases are being developed. They are potent analgesics but have fewer side effects than the opiates. List of Abbreviations: AD, Alzheimer’s disease; APN, aminopeptidase N; AVP, arginine‐vasopressin; ACE, angiotensin‐converting enzyme; CPE, carboxypeptidase E; CSF, cerebrospinal fluid; CM, conditioned medium; DAT, dementia of Alzheimer type; DAP, dipeptidyl aminopeptidase; Leu‐Enk, leucine‐enkephalin; Leu bNA, leucine b‐naphthylamide; MHC, major histocompatibility complex; Met‐Enk, methionine‐enkephalin; MSA, multiple system atrophy; bNA, b‐naphthylamide; NAP, neuron‐specific aminopeptidase; NPY, neuropeptide Y; NEP, neutral endopeptidase; NE, norepinephrine; PD, Parkinson’s disease; pLAP, placental leucine aminopeptidase; PE, proenkephalin; PC1 and PC2, prohormone convertase 1 and 2; POMC, proopiomelanocortin; PSA, puromycin‐sensitive aminopeptidase; CCK‐8S, sulfated cholecystokinin‐8; TRH, thyrotrophin‐releasing hormone; VIP, vasoactive intestinal peptide

1

Introduction

Neuropeptidases are defined as peptidases that are active in the brain or as enzymes that degrade neuropeptides, peptide neuromodulators, or peptide hormones found in the brain. Brain peptidases participate in the general metabolism of peptides (proteins). In addition, they are possibly involved in neurotransmission. The neuropeptidases that have been actively studied in the last 15 years are emphasized here. For the other peptidases, please refer to the author’s former review (Hui and Lajtha, 1983). Neuropeptides play important roles in mental and neurological diseases (Beal et al., 1986; Vecsei and Widerlov, 1988, 1990; Wahlestedt et al., 1989). They are widely distributed in the central nervous system (CNS) and peripheral nervous system, where they serve as neurotransmitters and neuromodulators. Deficiency of neuropeptides, for example, ACTH, can cause various neurological signs (Sato et al., 1991). The neurophysiological abnormalities include slow wave activity on electroencephalograms, delayed conduction velocity of the peripheral nerves, and low amplitude of muscle action potentials. Recent interest has focused on the role of neuropeptides in degenerative neurological diseases. Understanding this role could help define what is specifically vulnerable to the pathological processes, thus leading to improvement in diagnosis and therapy.

2

Abnormalities of Neuropeptides and Neuropeptidases

2.1 Neuropeptides in Degenerative Diseases In the CNS, neuropeptides are co‐stored with catecholamines, especially norepinephrine (NE). Their involvement in pathologies is characterized by a noradrenergic impairment (Martignoni et al., 1992). In Parkinson’s disease (PD), and in multiple system atrophy (MSA) as well, a central noradrenergic deficit has been demonstrated, and in dementia of Alzheimer type (DAT), impaired noradrenergic transmission has been found. PD, MSA, and DAT patients showed a significant reduction in cerebrospinal fluid (CSF) neuropeptide Y (NPY) and NE levels (Beal and Martin, 1986; Martignoni et al., 1992).

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In Huntington’s disease, there are reduced concentrations of substance P, methionine‐enkephalin (Met‐Enk), and cholecystokinin in the basal ganglia; in contrast, the concentrations of somatostatin and thyrotrophin‐releasing hormone (TRH) are increased (Sagar et al., 1984). The basal ganglia neurons in which somatostatin and NPY are co‐localized are selectively preserved. A marked increase of somatostatin has been reported in the striatum in Huntington’s chorea. In another study, neurotensin was increased in the pallidum, while in Parkinson’s disease no significant changes in neurotensin content were observed (Beal et al., 1986; Palacios et al., 1990). In Alzheimer’s disease (AD) and in dementia associated with Parkinson’s disease, concentrations of cortical somatostatin are reduced (Jankovic and Maric, 1987). NPY and corticotrophin‐releasing factor are also reduced in the cerebral cortex of patients with AD. The reduced cortical concentrations of somatostatin and NPY in the cerebral cortex of patients with AD reflect a loss of neurons or terminals in which these two peptides are co‐localized (Beal and Martin, 1986). Somatostatin concentration is reduced in the hippocampus and neocortex of patients dying with Alzheimer’s type dementia. The levels of TRH and gonadotrophin‐releasing hormone neuropeptides are significantly reduced in patients with senile dementia. But, no changes were found in the telencephalic neurotensin content in senile dementia of the Alzheimer type (Sagar et al., 1984). Receptors of the neuropeptides are also changed in neurological diseases. Marked reductions in the density of somatostatin binding sites were observed in the caudate and putamen of patients with Huntington’s chorea. However, these receptors were well preserved in the nucleus accumbens and in the ventral aspects of the anterior putamen. No alteration of somatostatin receptors was observed in other brain areas. These findings suggest that somatostatin receptors in the human striatum are markedly downregulated or are localized where a population of neurons is at risk. Mice with megencephaly due to brain cell hypertrophy exhibit neurological and motor disturbances with seizure‐like activity, and disturbances in the insulin‐like growth factor system as well (Petersson et al., 1999). Its enkephalin messenger RNA expression is upregulated in the dentate gyrus granular layer and in ventral cortices, but downregulated in the CA1 pyramidal layer (Petersson et al., 2000). Enkephalin‐like immunoreactivity is elevated in mossy fibers of the hippocampus and the ventral cortices. Cholecystokinin has region‐specific up‐ and downregulation in the hippocampal formation and increased levels in ventral cortical regions. Galanin and NPY expression are increased in several layers and interneurons of the hippocampal formation, and in ventral cortices as well. In contrast, galanin‐like immunoreactivity is reduced in nerve terminals in the forebrain. It is not clear whether the mainly increased peptide levels contribute to the excessive growth of the brain or represent a consequence of this growth and/or of the neurological and motor disturbances. Following intracerebral innoculation of prions in mice, NPY mRNA expression is specifically upregulated in CA3 pyramidal neurons, whereas its expression in hilar neurons remains unaltered. Neuropeptide alterations preceding neurological dysfunction and neuronal death play a possible role in prion diseases (Diez et al., 1996). NPY may regulate glutamate release at the Schaffer collateral‐CA1 synapses in scrapie‐infected mice.

2.2 Neuropeptides in Cerebrospinal Fluid In the lumbar CSF of patients with AD, high levels of neurotensin are detected (Martignoni et al., 1992). There is a good correlation between plasma and CSF arginine‐vasopressin (AVP) values in most patients with Parkinson’s disease, dementia, cerebrovascular disease, multiple sclerosis, or other, mostly peripheral, neurological disorders (Neuser et al., 1984; Sorensen et al., 1985; Cramer et al., 1988; Vecsei and Widerlov, 1988). Significantly higher CSF‐AVP values were found in patients with cerebrovascular disease, whereas lower CSF values were found in patients with dementia and Parkinson’s disease (Reid and Morton, 1982; Sundquist et al., 1983). However, CSF/plasma gradients in patients with dementia and Parkinson’s disease were decreased to about 0.30, compared with 0.98 in patients with peripheral neurological disorders. In hypoxic‐ischemic encephalopathy, a significant elevation of plasma b‐endorphin concentration was

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observed (Sankaran et al., 1984). The association of increased plasma b‐endorphin concentration is linked with ongoing hypoxemia.

2.3 Neuropeptides in Neurological Diseases Met‐Enk, but not leucine‐enkephalin (Leu‐Enk), inhibits neurological signs and markedly diminishes the occurrence and intensity of histological lesions in the brain, and in the cervical, thoracic, and lumbar spinal cord in the development of experimental allergic encephalomyelitis (EAE) (Jankovic and Maric, 1987). Met‐enk, a potent immunomodulator/regulator, is important in the pathogenesis and prevention of the inflammatory autoimmune disease. N‐Acetylaspartylglutamate (NAAG), a neuropeptide found in millimolar concentrations in brain, is localized in subpopulations of glutamatergic, cholinergic, GABAergic, and noradrenergic neuronal systems. NAAG is released upon depolarization by a Caþþ‐dependent process and is an agonist at mGluR3 receptors and an antagonist at N‐methyl‐D‐aspartate (NMDA) receptors. NAAG exerts neuroprotective effects in a neonatal rat model of hypoxia–ischemia (Cai et al., 2002). The effects are largely associated with activation of the mGlu2/3 receptor. NAAG and b‐NAAG protect against injury induced by NMDA and hypoxia in primary spinal cord cultures (Yourick et al., 2003). Vasoactive intestinal peptide (VIP), a neuropeptide with a potent Leu‐Enk anti‐inflammatory effect, protects from inflammatory disorders. In most neurodegenerative disorders, including multiple sclerosis, Parkinson’s disease, and Alzheimer’s disease, massive neuronal cell death occurs as a consequence of an uncontrolled inflammatory response. VIP also has a neuroprotective effect by inhibiting the production of microglia‐derived proinflammatory factors (tumor necrosis factor a, interleukin‐1b, nitric oxide). It prevents neuronal cell death following brain trauma by reducing the inflammatory response of neighboring microglia. VIP is thus a valuable neuroprotective agent for the treatment of pathologic conditions in the CNS where inflammation‐induced neurodegeneration occurs (Delgado and Ganea, 2003). TRH induced neurological improvement in 17 of the 23 patients with amyotrophic lateral sclerosis (ALS) but little or none in the other ALS patients or in patients with other neurological diseases (Congia et al., 1991). Neurotropic murine coronavirus MHV‐JHM (JHMV) causes encephalitis and paralytic‐demyelinating disease in susceptible strains of mice and rats (Congia et al., 1991). It is a good model for human demyelinating diseases such as multiple sclerosis. Intracerebral administration of b‐endorphin reduced the incidence of JHMV‐induced paralytic‐demyelinating disease in mice. Protection from the disease was accompanied by significantly reduced virus replication in the brain (Gilmore et al., 1993). The data suggest that b‐endorphin engages immune mechanisms of host resistance to JHMV infection to protect the mice from disease. Peptides derived from ACTH and MSH help post‐lesion repair mechanisms in the peripheral nervous system by enhancing the early sprouting response of the damaged nerve. These peptides prevent cisplatin neuropathy in women suffering from ovarian cancer. Treatment based on nonendocrine fragments of ACTH/MSH could be a therapeutic option in cisplatin neuropathy (Gispen et al., 1992).

2.4 Neuropeptide Degradation in Neurological Diseases Mutations in tripeptidyl peptidase (TPP‐I) have recently been associated with a lysosomal storage disease, late infantile neuronal ceroid lipofuscinosis (CLN2) (Tomkinson, 1999; Junaid et al., 2000; Golabek et al., 2003). This disease is characterized by the accumulation of proteinaceous and auto‐fluorescent material within the lysosomes of neurons, which undergo massive cell death during the course of the disease (Bernardini and Warburton, 2002; Warburton and Bernardini, 2002; Wujek et al., 2004). TPP‐I is required for the partial or complete digestion of certain neuropeptides by brain lysosomes. Dipeptidyl peptidase‐I expressed in other tissues has extensive activity on peptides and can compensate for the loss of TPP‐I (Bernardini and Warburton, 2002). The levels of NAAG and the activity of carboxypeptidase II are altered in a regionally specific fashion in several neuropsychiatric disorders (Coyle, 1997). In Alzheimer’s postmortem brain, somatostatin‐28

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degradation is increased in Brodmann area 22 whereas substance P degradation is increased in the temporal cortex. Changes in the degradation of these neuropeptides that are affected in AD correlate with alterations in the activity of specific neuropeptidases. Trypsin‐like serine protease activity is increased in Brodmann area 22 in AD. It parallels the increased degradation of somatostatin‐28. The decreased activity of neutral endopeptidase 24.15 (NEP; EC 3.4.24.15) in the temporal cortex corresponds to the decreased degradation of substance P (Waters and Davis, 1995, 1997).

2.5 Exopeptidases in Aging and in Neurological Diseases The process of aging involves alterations in the activity of peptidases and proteases, although the precise changes have not yet been fully characterized. The activity of the soluble fractions of prolyl endopeptidase was reduced in the lungs of aged animals (Waters and Davis, 1997). Reduced activity of soluble pyroglutamyl peptidase I and aminopeptidase N (APN) were detected in the aged kidney and heart, respectively. In contrast, increased activity of particulate prolyl endopeptidase was detected in the brain stem. Most of these changes can be correlated with known alterations in the levels of peptides controlled by each enzyme. Puromycin‐sensitive aminopeptidase (PSA) functions as a trimming enzyme in the major histocompatibility complex (MHC) class I pathway, which is activated in brains of patients with AD. In these brains, intensely stained cells were found to be rich in the cerebral cortex. Double immunofluorescence studies confirmed that PSA‐positive cells were reactive microglia. Such PSA‐positive reactive microglia tended to be located in and around senile plaques and were observed to be associated with neurons containing neurofibrillary tangles. The microglia PSA may be associated with the pathological conditions of AD (Minnasch et al., 2003). Alanyl aminopeptidase activity was lower in the CSF of patients with AD, whereas no differences in CSF were detected in regard to the remaining aminopeptidases (Montes et al., 1998). No changes were found in the levels of amino acids in CSF or plasma. The plasma/CSF ratio for aminopeptidase activities was higher in patients with AD, although the difference was significant only for alanyl aminopeptidase (Iribar et al., 1998). In brains of patients with sporadic AD, decreased neuronal expression of a brain‐specific carboxypeptidase B (CPB) and clusters of microglia with peptidase immunoreactivity associated with its extracellular deposition were detected (Matsumoto et al., 2000). Brain CPB has a physiological function in APP processing and may have significance in AD pathophysiology.

2.6 Endopeptidases in Neurological Diseases A DNA polymorphism at the angiotensin‐converting enzyme (ACE) gene has been linked to the risk for late onset Alzheimer’s disease (Alvarez et al., 1999). Increased frequency of the ACE‐I allele has been found in patients with AD. In the distribution of an insertion (I)/deletion (D) polymorphism of ACE in patients with AD, an association between AD and ACE genotypes or alleles was found. The frequency of II genotypes in AD was 1.4 times higher than in controls, while that of DD genotypes was only 0.4 times as high. The altered distribution of ACE alleles in patients appeared to be independent of apolipoprotein E (Hu et al., 1999). Conditioned medium (CM) of NEP 24.15 antisense‐transfected neuroblastoma has a significantly higher level of amyloid b (Ab) (Yamin et al., 1999). Furthermore, synthetic Ab‐degradation is increased or decreased following incubation with CM of sense‐ or antisense‐transfected cells, respectively. Soluble Ab1–42 is degraded more slowly than soluble Ab1–40, while aggregated Ab1–42 showed almost no degradation. Pretreatment of CM with serine proteinase inhibitors completely inhibits Ab degradation. Additionally, a serpin family inhibitor tightly associated with plaques and elevated in brains of patients with AD blocks Ab degradation. Recombinant NEP 24.15 alone does not degrade Ab. 14C‐Diisopropyl fluorophosphate‐radiolabeled CM from NEP‐overexpressing cells contains increased levels of several active serine proteinases suggesting that NEP activates one or more Ab‐degrading serine proteases. The serpin inhibitor

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causes Ab accumulation by inhibiting an Ab‐degrading enzyme or by direct binding to Ab, rendering it degradation resistant. Insulin‐degrading enzyme is the main soluble Ab‐degrading enzyme at neutral pH in the human brain. The highest Ab protein‐degrading activity in the soluble fraction occurs between pH 4 and 5, and this aspartyl protease is inhibited by pepstatin. Synaptic membranes have much lower Ab protein‐degrading activity than the soluble fraction. EDTA inhibits the degrading activity but inhibitors of NEP 24.11, ‐24.15, ‐ 24.16, ACE, aminopeptidases, and carboxypeptidases have little or no effect (McDermott and Gibson, 1997). A novel Znþþ‐dependent metalloprotease activity associated with a Golgi apparatus‐ and plasma membrane‐enriched fraction can degrade endogenous APP to generate Ab containing C‐terminal fragments. This protease generates amyloidogenic fragments of APP that can serve as precursors for Ab (Mok et al., 1997).

3

Classification of Exo‐ and Endopeptidases

Exo‐ and endopeptidases are involved in the breakdown of larger (>30 amino acids) peptides into smaller ( Figure 21-1). PE requires proteolytic processing at paired basic residue sites (Lys‐Arg, Arg‐Arg, and Lys‐Lys), and at monobasic arginine sites as well, to liberate active enkephalins.

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. Figure 21-1 Structure of proenkephalin (PE). PE contains Met‐enkephalin (M), Leu‐enkephalin (L), Met‐Enk‐Arg‐Gly‐Leu (O), and Met‐Enk‐Arg‐Phe (H). K represents lysine and R arginine

Processing of PE begins with removal of the NH2‐terminal signal peptide at the rough endoplasmic reticulum (RER) during protein translation (Steiner et al., 1992; Hook et al., 1994; Hook and Yasothotnsrikul, 1998; Seidah et al., 1998; Acher et al., 2002). The PE is routed through the RER and Golgi apparatus, where they are packaged into secretory vesicles. There, endopeptidases cleave at the Lys‐Arg and other paired basic residue sites (Lys‐Lys, Arg‐Arg, Arg‐Lys) that flank the enkephalins within the precursor. The resultant peptide intermediates contain basic residue extensions at the COOH‐ and/or NH2‐termini that are later removed by carboxypeptidase E/H (Fricker, 1991; Hook and Yasothotnsrikul, 1998) and Arg/ Lys aminopeptidase (Hook and Yasothotnsrikul, 1998; Yasothornsrikul et al., 1998) respectively. This multistep proteolytic pathway generates bioactive enkephalins (> Figure 21-2). . Figure 21-2 Processing of a model proneuropeptide. This model proneuropeptide contains one copy of the processed peptide neurotransmitter or hormone. Cleavage at the dibasic site occurs at its NH2 terminus, between the dibasic residues, or at the COOH terminus of the dibasic residue site, represented by arrows at positions 1, 2, or 3, respectively. Removal of basic residues at COOH‐ and NH2‐termini are carried out by carboxypeptidase E/H and Arg/Lys aminopeptidase, respectively

Four proteases consisting of the cysteine protease known as ‘‘prohormone thiol protease’’ (PTP) (Yasothornsrikul et al., 1999), the subtilisin‐like prohormone convertase 1 and 2 (PC1 and PC2) (Azaryan et al., 1992), and a 70‐kDa aspartyl protease (Azaryan et al., 1995c), were found using full‐length enkephalin precursor as substrate (Krieger and Hook, 1991). In chromaffin granules, PTP is the major, protease having PE‐cleaving activity, PC1 and PC2 have comparably lower activities, and the 70‐kDa aspartyl protease has the least PE‐cleaving activity (Hook and Eiden, 1985; Azaryan et al., 1995c; Hook et al., 1996). PTP shows preference for processing PE, with minimal processing of POMC in vitro (Azaryan et al., 1995b; Schiller et al., 1996).

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4.1 Prohormone Thiol Protease PTP generates PE products in vitro that resemble those in vivo (Ungar and Phillips, 1983; Schiller et al., 1995; Hook et al., 1997). PTP has cleavage specificity for dibasic and monobasic processing sites that are cleaved within enkephalin‐containing peptide substrates. Native PTP cleaves PE and enkephalin‐containing peptide substrates at paired basic residue cleavage sites, and at monobasic Arg sites as well. The cleavage specificities of PTP for cleavage between and at the NH2‐terminal side of the dibasic residue sites indicate that resultant peptide intermediates possess basic residue extensions at their NH2‐termini (Loh et al., 1985; Azaryan and Hook, 1994a, b; Azaryan et al., 1995a). Studies of PE‐derived peptides demonstrate the presence of enkephalin peptides with basic residue extensions at their NH2‐termini (Hook and Eiden, 1984; Kreiger et al., 1992). These results indicate the necessity for an aminopeptidase that removes Arg and Lys residues from NH2‐termini of peptide intermediates as one of the later steps in PE processing. PTP has not been fully purified and characterized because of the trace yield. It was partially purified from bovine medullary chromaffin granules. The soluble protease is a glycoprotein with a pI of 6.0 and a pH optimum of 5.5 (Krieger and Hook, 1991). It is a thiol enzyme as shown by its dependence of dithiothreitol and inhibition by p‐hydroxymercuribenzoate, mercuric chloride, cystatin C, and E‐64. It was reported that PTP possesses a unique NH2‐terminal primary sequence, which is not homologous to other known proteases (Tezaosudusm et al., 1995). However, further work is needed to show that PTP is a novel enzyme. Recently, Yasothornsrikul et al. reported that secretory vesicle cathepsin L is the responsible cysteine protease of chromaffin granules for converting PE to the active enkephalin peptide neurotransmitter (Yasothornsrikul et al., 2003). The cathepsin L activity was identified by affinity labeling with an activity‐ based probe for cysteine proteases followed by mass spectrometry for peptide sequencing. Production of Met‐Enk by cathepsin L occurred by proteolytic processing at dibasic and monobasic prohormone‐processing sites. Co‐localization of cathepsin L with Met‐Enk in secretory vesicles of neuroendocrine chromaffin cells was shown by immunofluorescent confocal and immunoelectron microscopy. Cathepsin L was co‐secreted with Met‐Enk. In cathepsin L‐gene‐knockout mice, significant reduction in Met‐Enk levels in brain occurred with an increase in the relative amounts of enkephalin precursor (Yasothornsrikul et al., 2003).

4.2 Prohormone Convertase 1 and 2 PC1 and PC2 are proprotein convertase members of the mammalian subtilisin‐like family. They cleave proinsulin and other prohormones primarily at the COOH‐terminal side of paired basic residues, with some cleavage at the NH2‐terminal side of a single arginine residue of a peptide (Hwang et al., 2000). PTP cleaves at the NH2‐terminal side of paired basic residues and between the two basic residues, whereas PC1 and PC2 cleave pro‐opiomelanocortin (POMC) between the two basic residues and at the COOH‐terminal side of the dibasic residues. The proteases have highly conserved primary sequences with respect to signal sequence, pro‐segment, catalytic domain, and P domain. The mammalian PC1 and PC2 contain catalytic triad residues Asp, His, and Ser. The bovine PC1 contains Asn as the oxyanion hole residue and PC2 contains Asp as the oxyanion hole residue. Each of them possesses the P domain with a functional RRGDL motif (Hwang et al., 2000).

4.3 Carboxypeptidase H/E The cleavage specificities of the processing enzymes require carboxypeptidase E/H and aminopeptidase for the removal of NH2‐ and COOH‐terminal basic residues from the peptide intermediates (Dhanvantari et al., 2002; Wei et al., 2003). Carboxypeptidase E (CPE) (EC 3.4.17.10; carboxypeptidase H) removes basic amino acids from the COOH terminus of peptides to make them biologically active. A variety of neuropeptide‐processing endopeptidases cleave at specific cleavage sites, at the COOH ends of the basic residues, generating intermediates with the COOH‐terminal amino acids. However, a single carboxypeptidase has been implicated in the processing of mammalian neuropeptides (Wei et al., 2002).

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CPE is present within secretory granules in both a soluble form and a form that is membrane bound at pH 5.5 but soluble at neutral pH (Fricker et al., 1990). Combined polar and hydrophobic interactions of the COOH‐terminal peptide appear to be responsible for the reversible pH‐dependent association of CPE with membranes (Silva et al., 1995; Dhanvantari et al., 2002).

4.4 Arg/Lys Aminopeptidase Arg0‐Met‐Enk was found and isolated from bovine striatum (Hui et al., 1994). The structure of the purified material was identified by microsequencing and mass spectrometry as the hexapeptide Arg‐Tyr‐ Gly‐Gly‐Phe‐Met. Ninety percent of the purified peptide was Arg0‐[O]Met‐Enk, consisting of equimolar Met(R)‐ and (S)‐sulfoxide. The existence of this enkephalin intermediate indicates that PTP is the putative processing enzyme. It will be interesting if there is an enzyme that specifically removes the N‐terminal basic amino acid to turn on enkephalin activity. There is an aminopeptidase activity in neurosecretory vesicles that converts Arg0‐Met‐Enk to Met‐Enk (Gainer et al., 1984; Hook and Eiden, 1984). Both arginine aminopeptidase (RAP) and lysine aminopeptidase (KAP) activities were found in neurosecretory vesicles of chromaffin granules (Yasothornsrikul et al., 1998). They are involved with reduced cysteinyl residues. The majority of the RAP/KAP activity resides with the soluble component of chromaffin granules, rather than the membrane component. RAP, but not KAP, is stimulated with NaCl. The RAP and KAP activities have pH optima at 6.7 and 7.0, respectively. RAP has a lower Km. Both enzymes, possible metalloproteases, are inhibited by the specific aminopeptidase inhibitors bestatin, amastatin, and arphamenine. KAP activity is partially inhibited by Niþþ and Znþþ, whereas RAP activity is affected less. The chromaffin granule RAP/KAP resembles rat aminopeptidase B, which specifically removes basic residues from NH2‐termini of peptides. Recently we found that KAP is different from RAP (Hui and Hui, unpublished observations). KAP and RAP can be physically separated by FPLC with Mono Q where KAP is eluted with less NaCl. The molecular weight of KAP was determined by gel‐filtration to be 62,000 daltons, which is 10,000 more than RAP. Using aminoacyl b‐naphthylamides (bNA) as substrate, KAP prefers lysine five times more than arginine, but RAP prefers arginine one time more than lysine. NaCl can activate KAP. RAP is inhibited by thiol blocking agents and is most sensitive to arphamenine B. In contrast, KAP is most sensitive to bestatin. Leukocyte‐derived RAP is a 960‐amino‐acid protein with significant homology to placental leucine aminopeptidase (pLAP) and adipocyte‐derived leucine aminopeptidase (aLAP/ERAP1) (Tanioka et al., 2003; Bolumar et al., 2003). It contains the HEXXH(X)18E zinc‐binding motif, a characteristic of the M1 family of zinc metallopeptidases. It is a subfamily with pLAP and aLAP/ERAP1 in the M1 family. L‐RAP located in the lumenal side of the endoplasmic reticulum and has a preference for arginine with synthetic substrates. Its substrate specificity is restricted. It cleaves angiotensin III, kallidin, and the N‐terminal extended precursors of MHC class I‐presented antigenic peptides.

5

Enkephalin‐Degrading Enzymes

Study of neurotransmitter enzymes has made a substantial contribution to our understanding of synaptic biochemistry. Some of their inhibitors turn out to be valuable therapeutics. Enkephalin binds to the opiate receptors on the neuronal membrane to trigger intracellular functions. Its action is apparently terminated by synaptic degradation, since no uptake, internalization mechanism, N‐acetylation, O‐sulfation, phosphorylation, or glycosylation has been found (Patey et al., 1981; Goodman et al., 1983, Schwartz et al., 1985). In the CNS, several sets of peptidases are capable of cleaving enkephalins (Tye‐Gly‐Gly‐Phe‐Met [Leu]) at different sites: aminopeptidases (APN, PSA, neuron‐specific aminopeptidase (NAP)) at the Tyr‐ Gly amide bond, dipeptidyl aminopeptidase (DAP) at the Gly‐Gly bond, carboxylpeptidase at the Phe‐Met bond, and ACE and NEP, both at the Gly‐Phe bond (> Figure 21-3).

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. Figure 21-3 Degradation of enkephalin by peptidases at different sites. NAP, neuron‐specific aminopeptidase; APN, aminopeptidase N; PSA, puromycin‐sensitive aminopeptidase; DAP, dipeptidyl aminopeptidase; ACE, angiotensin‐ converting endopeptidase; NEP, endopeptidase 24.11

5.1 Endopeptidase 24.11 Endopeptidase 24.11 (NEP 24.11) (neprilysin; EC 3.4.24.11) participates in the postsecretory processing and metabolism of synaptically released neuropeptides (Turner et al., 1996). Inhibition of NEP by thiorphan (Patey et al., 1981) induces analgesia (Llorens et al., 1980), but it has been controversial whether it is neuronal (Horsthemke et al., 1983; Barnes et al., 1988). NEP 24.11 is a cell‐surface zinc metallopeptidase with specificity directed toward the hydrolysis of peptide bonds at the N‐terminal side of amino acids of hydrophobic residues: Phe, Tyr, Leu, and Trp (Turner and Barnes, 1995; Hooper and Turner, 1988). NEP 24.11 is sensitive to the chelating agents EDTA and 1,10‐phenanthroline. It is highly susceptible to phosphoramidon (Ki ¼ 2 nM) and thiorphan. The enzyme is distributed widely and in the CNS serves to inactivate enkephalins, tachykinins, and somatostatin (Turner et al., 1996). In vitro, the enzyme hydrolyzes neutrophil chemotactic peptide, atrial natriuretic peptides, vasoactive intestinal polypeptide, and calcitonin gene‐related peptide. NEP 24.11 was found to be co‐localized on the plasma membrane of substance P‐rich boutons in the substantia nigra, suggesting its probable role as a neuropeptidase (Turner et al., 2001). Molecular cloning of NEP 24.11 revealed its identity with the common acute lymphoblastic leukemia antigen (CD10) (Howell et al., 1991), implying its primary role in the immune system. Nonetheless, this underscores the similarities of signal mechanisms in, and interaction between, the immune and nervous systems.

5.2 Angiotensin‐Converting Enzyme ACE (EC 3.4.15.1, peptidyl dipeptidase A), a peptidyl carboxypeptidase, is responsible for the conversion of angiotensin I to the potent vasoconstrictor angiotensin II and for the degradation of bradykinin (Chai and Mendelsohn, 1995). It hydrolyzes a range of peptides including opioid peptides, neurotensin, bombesin, tachykinins, and luteinizing hormone‐releasing hormone (Chai and Mendelsohn, 1995). ACE is a membrane‐bound ectoenzyme of 146 kDa that consists of a large extracellular domain with a transmembrane anchor and a small intracellular carboxyl terminus (Soubrier et al., 1988). Its extracellular domain contains two regions of high sequence homology, each containing an active site sequence. Although the two sites are catalytically active, the C‐terminal one is more responsible for most of the hydrolysis of angiotensin I and for the binding of inhibitors (Wei et al., 1991; Perich et al., 1992). The N‐terminal site has structural constraints that limit the binding of some substrates and drugs (Wei et al., 1992). In human beings, ACE has two forms and two functions. Testicular ACE (tACE) has only one active site and has a sequence identical to the C terminus of the somatic ACE. Somatic ACE exists in most cells, and tACE, which is half the size of somatic ACE, is found only in the testis (Ehlers and Riordan, 1991; Perich et al., 1992). The enzyme is composed of a‐helices for the most part, and incorporates a zinc ion and two chloride ions (Schullenk and Wilson, 1988). Chloride ions activate tACE, and they interact with the substrate as well. However, the structure of tACE places the chloride ions outside the active site. Therefore, they play an

Neuropeptidases

21

indirect role in substrate activation (Wei et al., 1991). The zinc ion lies in the active site and interacts directly with inhibitor lisinopril (Wei et al., 1992). The structure of tACE shows it to be similar to ACE forms found in other species, and in other zinc‐containing metallopeptidases as well. Testicular ACE is roughly an ellipsoid in shape, divided into two subdomains by a central groove. The active site is toward the bottom of the groove and is capped by an N‐terminal lid, which prevents large molecules from fitting into the active site. Somatic ACE consists of two parts (called the N‐ and C‐domains), each with a different function. The newly described tACE structure, which is identical to the C‐domain structure of somatic ACE, now serves as a template for the search of specific domain‐selective ACE inhibitors (Wei et al., 1991).

5.3 Aminopeptidases The liberation of Tyr is the major mode of enkephalin inactivation with intact‐cell and cell‐free preparations (Hui and Lajtha, 1983; Lentzen and Palenker, 1983). The aminopeptidase inhibitors exert a dose‐ dependent, naloxone‐reversible, analgesic effect when administered to mice (Zhang et al., 1982; De La Baume et al., 1983; Herman et al., 1985). In addition, aminopeptidases that are sensitive to puromycin underlie a variety of specialized CNS functions: memory (Herman et al., 1985), amnesia (Eisenstein et al., 1983), apoptosis (Tobler et al., 1997), and schizophrenia (Hui et al., 1995). Brain aminopeptidase activity can be simply classified into PSA (gene)‐dependent; PSA‐independent; but puromycin‐sensitive, and puromycin‐insensitive. Earlier, it had been suggested that enkephalin binding to the opiate receptor was coupled to subsequent aminopeptidase degradation (Knight and Klee, 1978). APN was copurified with opiate receptors (Hui et al., 1985), though APN was later found localized exclusively in the blood microvessels (Solhonne et al., 1987; McLellan et al., 1988). More than 20 aminopeptidases (EC 3.4.11) have the capability to liberate various amino acid residues from the NH2‐termini of peptide substrates. These ubiquitous enzymes are mostly concentrated in brain and kidney. They are classified according to the preference for the NH2‐terminal amino acid of the substrates, their location, the susceptibility to inhibitors, the metal ion content, the residues that link the metal to the enzyme, and the pH for maximal activity. These enzymes are considered to have their original structures, substrate specificities, and specific locations depending on their physiological roles.

5.3.1 Aminopeptidase N Leu‐Enk is readily hydrolyzed to free tyrosine and Gly‐Gly‐Phe‐Leu by APN on the surface of microglia (Lucius et al., 1995). APN activity in microglia is higher than in rat peripheral monocytes and macrophages. Sequence data has shown its strong homology to CD13, a 150‐kDa cell‐surface glycoprotein (Look et al., 1989; Razak and Newland, 1992). Its nucleotide sequence predicts a 967‐amino‐acid integral membrane protein with a single, 24‐amino‐acid hydrophobic segment near the amino terminus. Amino‐terminal protein sequence analysis of CD13 molecules indicated that the hydrophobic segment is not cleaved, but rather serves both as a signal for membrane insertion and as a stable membrane‐spanning segment. The remainder of the molecule consists of a large extracellular carboxy‐terminal domain, which contains a pentapeptide consensus sequence characteristic of members of the zinc‐binding metalloprotease superfamily. APN, a membrane‐bound glycoprotein, is involved in the metabolism of regulatory peptides by diverse cell types, including small intestinal and renal tubular epithelial cells, macrophages, granulocytes, and synaptic membranes prepared from cells of the CNS (Look et al., 1989).

5.3.2 Puromycin‐Sensitive Aminopeptidase The concentration of PSA is highest in the brain (McLellan et al., 1988), 100‐fold that of APN (Solhonne et al., 1987). Eighty percent of the brain PSA is cytosolic. Using Ala‐b‐naphthylamide as substrate for rat brain PSA, Vmax was shown to be pH independent over the range of 5.5–9.0, while the Km exhibited a pKa

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of 7.7. This latter value corresponds to the pKa of the amino group of the substrate. Using X‐Ala and X‐Leu to examine the specificity of the P1 site, it was found that Arg and Lys exhibit the highest affinity, followed by Met, Val, Leu, Trp, and Phe, which bind congruently 5‐ to 20‐fold less. Although the Km varied more than 20‐fold within this series, Vmax showed considerably less variation. Significantly weaker binding was observed with P1 Gly, Ala, Ser, or Pro, with no binding detectable with P1 Glu. The presence of P0 1 Leu compared to P0 1 Ala results in an approximate tenfold decrease in Km with little change in Vmax. The effect of varying P0 1 residues was examined with the series Leu‐X. In this case, basic and hydrophobic amino acids, with the exception of Val, all exhibit nearly the same Km. The binding of Arg‐Arg and Lys‐Lys showed the same Km as obtained for Arg‐Leu or Lys‐Leu, respectively. When Leu‐Ser‐Phe was compared with Leu‐Ser, the P0 2 residue led to a 100‐fold decrease in Km and slightly less than a fivefold increase in Vmax. In contrast, the addition of a P0 2 Met to Leu‐Trp resulted in only a threefold decrease in Km and a threefold increase in Vmax. PSA prefers a basic or hydrophobic residue in the P1 and P0 1 sites and the subsite–subsite interactions primarily affect binding (Johnson and Hersh, 1990). Brain has soluble and membrane‐associated forms of PSA. All tissues contained significant levels of the soluble enzyme form, with this enzyme accounting for greater than 90% of the arylamidase activity in brain, heart, and skeletal muscle. In contrast to the results obtained with the soluble enzyme form, brain was the only tissue found to contain the membrane‐associated enzyme form. The brain membrane‐associated enzyme could be distinguished from the membrane‐associated aminopeptidase activity in other tissues on the basis of its sensitivity to inhibition by puromycin (McLellan et al., 1988). Each membrane‐bound and cytosolic PSA is composed of a single polypeptide of a molecular mass of 100 kDa. The anti‐soluble aminopeptidase antiserum reacts with both enzyme forms on immunoblots and inhibits both with nearly identical inhibition curves. The isoelectric points (pI ¼ 5.0) of the two forms were shown to be identical. N‐terminal sequencing yielded a common sequence (P‐E‐K‐R‐P‐F‐E‐R‐L‐P‐T‐ E‐V‐S‐P‐I‐N‐Y) for the two enzyme forms, and peptide mapping yielded 26 peptides that also appeared to be identical between the two enzyme forms. The membrane form of the PSA is identical to the soluble enzyme; it associates with the membrane by interactions with other integral membrane proteins (Dyer et al., 1990). A membrane‐bound aminopeptidase was purified from the rat brain (Hui et al., 1983c). The enzyme was extracted with 1% Triton X‐100 and purified by chromatography successively on DEAE‐Sepharose CL‐6B, Bio‐Gel HTP, and Sephadex G‐200 columns. The purified enzyme showed one band on disc gel electrophoresis and two bands on sodium dodecyl sulfate (SDS) electrophoresis with molecular weights of 62,000 and 66,000. The aminopeptidase has a pH optimum of 7.0, a Km of 0.28 mM, and a Vmax of 45 mmol/mg of protein/min for Met‐Enk. It releases tyrosine from Met‐Enk, but it does not split the byproduct, GlyGlyPheMet. It hydrolyzes neutral and basic aminoacyl bNA, but not g‐ or b‐endorphin, or dynorphin. The enzyme is inhibited by the specific aminopeptidase inhibitors amastatin, bestatin, and bestatin‐Gly. Its subcellular localization, substrate specificity, pH optimum, and molecular weight distinguish it from leucine aminopeptidase, aminopeptidase A, aminopeptidase B, aminopeptidase M, and the soluble aminopeptidase for enkephalin degradation. PSA is inhibited by enkephalin‐containing polypeptides derived from proenkephalin A, proenkephalin B, and proopiomelanocortin (Hui et al., 1983a). Of the peptides, Arg0‐Met‐Enk was the most potent inhibitor with an IC50 of about 0.6 mM; it was more effective than bestatin. This inhibition was partly due to substrate competition. PSA hydrolyzed Arg0‐Met‐Enk to Arg, Tyr, and Gly‐Gly‐Phe‐Met in a substrate‐ inhibited manner. The hexapeptide also inhibited the breakdown of Arg‐ and Tyr‐bNA by the membrane PSA. Since Arg0‐Met‐Enk did not inhibit leucine aminopeptidase, it was a more selective inhibitor than bestatin of Met‐Enk breakdown by aminopeptidases. Arg0‐Met‐Enk also inhibited enkephalin breakdown by synaptosomal plasma membranes but not by brain slices. PSA Gene The human PSA gene is composed of 23 exons and 22 introns and spans approximately 40 kb of chromosome 17 at the interval 17q12–21 (Thompson et al., 1999). The gene (NPEPPS) was physically mapped to q21.2!q21.32 of chromosome 17 using fluorescence in situ hybridization (Bauer et al., 2001). PSA is 27–40% homologous to several known Znþþ‐binding aminopeptidases including APN

Neuropeptidases

21

(Constam et al., 1995). An analysis of the 50 ‐end of the human PSA transcript reveals that the translational start site corresponds to nt 210 of the human PSA cDNA. A comparison of the exon/exon boundaries of the human PSA gene with those of the human APN gene shows little conservation, suggesting that the two genes, which are closely related in protein sequence, diverged early during evolution (Thompson et al., 1999). The tissue distribution of PSA is a polymorphism within the coding region and the complete 30 ‐UTR (Bauer et al., 2001). Putative catalytic residues of PSA, Cys146, Glu338, and Lys396, were mutated and the resultant mutant enzymes ApPS C146S exhibited normal catalytic activity (Thompson and Hersh, 2003). ApPS E338A exhibited decreased substrate binding, and ApPS K396I exhibited decreases in both substrate binding and catalysis. ApPS K396I and ApPS Y394F were analyzed with respect to transition‐ state inhibitor binding. No effect was seen with the K396I mutation, but ApPS Y394F exhibited a 3.3‐ fold lower affinity for RB‐3014, a transition‐state inhibitor. Thus, Tyr394 is involved in transition‐state stabilization. Conversion of glutamate 309 to glutamine resulted in a 5,000‐ to 15,000‐fold reduction in catalytic activity (Thompson et al., 2003). Conversion of this residue to alanine caused a 25,000‐ to 100,000‐fold decrease in activity, while the glutamate to valine mutation was the most dramatic, reducing catalytic activity 300,000‐ to 500,000‐fold. In contrast to the dramatic effect on catalysis, all three mutations produced relatively small (1.5‐ to 4‐fold) effects on substrate binding affinity. Mutation of a conserved tyrosine, Y394, to phenylalanine resulted in a 1,000‐fold decrease in kcat, with little effect on binding. Glutamate 309 acts as a general acid/base catalyst. Its mutation E309V converts the enzyme into an inactive binding protein. The effect of mutating tyrosine 394 is consistent with involvement of this residue in transition‐state stabilization (Thompson et al., 2003). Homozygous goku mice generated by gene‐trap mutation showed dwarfism, a marked increase in anxiety, and an analgesic effect (Osada et al., 1999). The function of PSA is disrupted in transcriptional arrest of the PSA gene and a drastic decrease of aminopeptidase activity. Because the PSA gene is strongly expressed in the brain, especially in the striatum and hippocampus, the PSA gene is required for normal growth and for behavior associated with anxiety and pain (Osada et al., 1999). PSA Functions In patients with schizophrenia, prefrontal cingulate and frontal cortices, thalamus, hippocampus, hypothalamus, and outer globus pallidus contained significantly less PSA as quantified by Western blot analysis than the corresponding areas from control subjects (Hui et al., 1995). Aminopeptidases may play an important role in the processes of tolerance and withdrawal associated with morphine administration. Increased activity of PSA was found in the brain cortex of heroin addicts in humans (Larrinaga et al., 2005). In rats treated with morphine, the activity of soluble PSA was found to be higher in the frontal cortex (Irazusta et al., 2003). In contrast, rats experiencing withdrawal symptoms presented decreased levels of aminopeptidase activity in certain brain areas. The activity of APN in the hippocampus and soluble PSA in the frontal cortex were lower in rats experiencing naloxone‐precipitated withdrawal symptoms. However, the activity of the aminopeptidases in vitro was unaltered by incubation with morphine, suggesting an indirect action of this opioid upon the aminopeptidases. PSA participates in proteolytic events essential for cell growth and viability (Constam et al., 1995). Proteolysis involves a cascade of enzymes including 26S proteasome. PSA is localized to the cytoplasm and to the nucleus and is associated with microtubules of the spindle apparatus during mitosis. Puromycin and bestatin both arrested the cell cycle, leading to an accumulation of cells in the G2/M phase, and ultimately induced cells to undergo apoptosis at concentrations that inhibit PSA. PSA in Development PSA activity increases twofold in the synaptosomal and mitochondrial fractions during the period of axonal and dendritic growth. This enzyme also has significant age‐related changes in the nuclear fraction. Significant developmental changes of APN are found only in the myelinic and microsomal fractions and they are less significant than those found for PSA (de Gandarias et al., 1999). PSA is mainly transported by anterograde axonal flow and plays a role in the metabolism of neuropeptides in nerve terminals or synaptic clefts (Yamamoto et al., 2002).

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5.3.3 Neuron‐Specific Aminopeptidases Strong aminopeptidase activity has been found in the CNS neuronal cell bodies by using histochemistry and in situ hybridization (Constam et al., 1995). Synapses, channels of communication between neurons, are sites of storage of information that is encoded by genes and by experience—memories. The nerve terminal is a site of integration where the signals it receives can modify the secretory response to an action potential. The release of neuropeptides (neurotransmitters) is the final response of a nerve to the excitatory and inhibitory inputs that converge upon it. Because it is the site of final signal output, the nerve terminal is an especially sensitive and critical point of control for neurons. To understand how these modifications occur by studying the neuropeptide metabolism in nerve terminals presents a neural anchor to decipher how the brain works. If anatomical distribution (specific location) associated with limited substrate specificity does constitute a ‘‘functional’’ specificity, it would be interesting to find an aminopeptidase specific for the CNS synapse or neuron (Shaw and cook 1978). Knowledge of specific enkephalin‐ inactivating aminopeptidases will facilitate our understanding on how the neuropeptide functions at the synapses. The widely distributed aminopeptidases in brain possess overlapping substrate specificity. In order to quantify accurately an aminopeptidase in biological samples, a post‐column continuous‐flow aminopeptidase detector was developed (Hui and Hui, 1996). Its conjugation with an FPLC provides a fast, sensitive, specific, and reliable method for brain aminopeptidase screening and quantitation. An enzyme of interest is separated from interfering peptidases, activators, and endogenous inhibitors in the samples (Hui and Hui, 1996). Using leucine b‐naphthylamide (Leu bNA) as substrate, two novel neuron‐specific enkephalin‐degrading aminopeptidases (NAP1 and NAP2) are separated from PSA and from each other with a FPLC Mono Q (> Figure 21-4). They are present only in the mammalian CNS and not in peripheral tissues, serum, or sciatic nerve (> Table 21-1). The two aminopeptidases are present exclusively in neurons, not in other nerve cells and neuroblastomas. The regional distribution of NAP1 and NAP2 is different. The highest NAP1 was found in the hippocampus, whereas the highest NAP2 was in the colliculus. Hypothalamus has the highest ratio of NAP1:NAP2. Both aminopeptidases are enriched in the synaptosomes with NAP1 > NAP2. . Figure 21-4 Zymogram of aminopeptidases by FPLC‐aminopeptidase analyzer

In rat brains, 85% of NAP1 is soluble and the rest is associated with membranes despite their relationship not being established (Hui et al., 1998). It is higher in the synaptosomes, of which the lysate has a specific activity 350% of that of the cytosol (S2) fraction, and 200% that of the membrane fraction. In rat cerebrocortical neuron cultures, its activity is 33% of the total aminopeptidase activity; in cerebellar granule cells, it is 12%, and it is absent in astrocytes. NAP1 cannot be found in glioma C6 and neuroblastoma

21

Neuropeptidases . Table 21-1 Distribution of NAPs and PSA in different rat tissues Activity (units*)

CNS Whole brain Brian regions: cortex, striatum, corpus callosum, hypothalamus, midbrain, hippocampus, cerebellum, medulla oblongata, superior and inferior colliculus Spinal cord (cervical, sacral) Olfactory bulb Other tissues Kidney, spleen, heart, intestinal mucosa, skeletal muscle, testis, liver, adrenal gland, pituitary, sciatic nerve, retina Serum

NAP1

NAP2

PSA

82 40–123

40 14–58

400 140–430

32–40 21

16–30 7.8

141–165 230

n.d.

n.d.

34–250

n.d

n.d.

2.5

50 ml of the S3 fraction of 2.5 mg of tissue or serum was submitted to the automatic FPLC‐aminopeptidase analyzer *arbitrary fluorescence units n.d.: not detectable

SK‐N‐SH cells. Its predominance in brain synaptosomes suggests that NAP1 plays a significant role in neurotransmission and synaptic differentiation. Purification and Characterization of NAP NAP1 was purified from rat brain to homogeneity by ammonium sulfate fractionation, followed by column chromatography, successively on phenyl‐Sepharose, Sephadex G‐200, and twice on Mono Q FPLC (Hui et al., 1998). The purified single‐chain enzyme was estimated to be 110 kDa. It has a pI of 5.25 and a pH optimum of 7.0. Only Mgþþ restores the activity of the apoenzyme. The neutral aminopeptidase hydrolyzes bNA of amino acids with aliphatic, polar uncharged, positively charged, or aromatic side chains. It has a Km of 95 mM and a kcat of 7.8 s1 on Met‐enk, releasing only the N‐terminal tyrosine. The thiol‐dependent metalloenzyme is most sensitive to amastatin inhibition with a Ki of 0.04 mM and is the aminopeptidase most sensitive to puromycin. Its properties are different from those of the ubiquitous PSA obtained from the same enzyme preparation. The blocked N terminus, substrate and inhibitor specificity, hydrolytic coefficiency, metal effects, pI, molecular weight, and catalytic site show that this enzyme is distinct from all other known aminopeptidases (Hui et al., 1998). Using an anti‐PSA IgG to screen a rat brain cDNA expression library a 1,561‐bp cDNA was isolated. Probing with this cDNA, we cloned a candidate 1,404‐bp cDNA (63.2% identity to mouse PSA) encoding the N‐terminal section of neural aminopeptidase. The nucleotide segment position at 875–1,404 is homologous to position 1,087–1,613 of PSA (96.4% identity), containing a sequence encoding a divalent metal‐binding motif, HEXXH(X)18E, of aminopeptidases (Shannon et al., 1989). The sequence at the 50 ‐end 1–874, with an identity 44.9% to PSA, is novel. It is void of the sequence encoding a universal N‐terminal PENKRPFERLPTEVSPINY of PSA (Dyer et al., 1990; Constam et al., 1995; Tobler et al., 1997). The blocked N‐terminal residue, though, waits to be identified. The cloning data imply that the purified enzyme is a unique aminopeptidase and a possible member of the PSA superfamily. NAP1 During Neuron Growth NAP1 was found in the rat hippocampus in all ages (Hui and Hui, 2003). It was lower in immature rat; the 19th embryonic‐day fetus contained the least. NAP1 increased steeply during the prenatal through the early post‐natal period, nine‐fold by the first month. The rate of increase diminished subsequently, increasing 20% in the second month and 13% in the third. The age‐dependent increase in NAP1 activity was parallel to its protein expression. The specific hydrolytic activity/NAP1 antigenicity in newborn, 15‐day‐old, and 30‐day‐old were 1.00, 0.88, and 1.00, respectively. Its growth profile was distinct from that of PSA. A similar difference between them was also found in the developing primary cerebellar granule cells. Puromycin (1–5 mM) blocked neurite outgrowth and caused apoptosis by non‐antibiotic effects.

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NAP1 Is a Putative Synaptic Enzyme If NAP1 is a synaptic enzyme, it could be released by electrical stimulation or inhibited by synaptic factors. Using a modified aminopeptidase analyzer (> Figure 21-5), we found a NAP1 inhibitor released by electric stimulation from slices of brain, but not in kidney or liver. Five hippocampal (or other tissue) slices (1 mg, 450 mm thick) were rinsed, equilibrated, and perfused with Krebs solution at 0.1 ml/min in a microvolume (0.5 ml) glass chamber. The perfusion effluent was mixed with NAP1 (> Figure 21-5) at a flow rate of 0.05 ml/min, and joined with Leu bNA (0.15 ml/min). The enzyme reaction proceeded in a delaying coil (37 C) for 3 min before the detection of the released . Figure 21-5 Flow diagram of the automated analyzer for NAP inhibitor released from tissue slices

b‐naphthylamide by fluorescence. The inhibitor activity was monitored continuously by measuring the decrease of free b‐naphthylamide. The inhibitor was released by electric stimulation at optimal conditions for neuropeptide release (20V, 20 Hz, 20ms, 300 shocks) (Milusheva et al., 1992). > Figure 21-6 shows a typical tracing of the NAP1 activity inhibited by electric stimuli on brain slices. With each stimulus, a 4‐min inhibitory effect, independent of its intensity (10–40 V) and frequency (10–40 Hz), was observed. The potency of the released inhibitor by a stimulus was equivalent to 1 nmol of bestatin. The potent inhibition was not due to substrate (100 mM Leu bNA) competition. It was due neither to serotonin, catecholamines, acetylcholine, amino acids, nor Met‐Enk, which had no inhibitory effect on NAP1. The perfusate showed no effect

. Figure 21-6 Synaptic aminopeptidase activity inhibited by factors released by electric stimulation (S) from hippocampal slices

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on radioreceptor bindings at d, m, or k sites indicating that the inhibition was not by enkephalins or enkephalin‐like peptides. The ninhydrin‐positive inhibitor could be retained on a reverse‐phase C18 Sep‐Pak column and eluted by methanol. Its activity disappeared after treatment with 6N HCl at 110 C for 16 hr. Our data indicate that the small‐peptide inhibitor was vesiculated in synapses. The inhibitor(s) was estimated to be 1,000 daltons by Sephadex G‐10.

6

Other Peptidases

6.1 Tripeptidyl Peptidase (TPP) Protein degradation is essential for the life and death of every cell. Proteins are broken down to their constitutive amino acids by a succession of peptidases, both in lysosomes and in the cytosol. TPP‐I and TPP‐II are enzymes that can ‘‘count to three’’ and release N‐terminal tripeptides from oligopeptides generated by different endopeptidases. The tripeptides are then degraded by other exopeptidases to release amino acids and dipeptides. The molecular weight of TPP‐I was calculated to be 280,000 and 290,000 by non‐denaturing polyacrylamide electrophoresis (PAGE) and gel filtration respectively, and to be 43,000 and 46,000 on SDS‐PAGE in the absence and presence of b‐mercaptoethanal, respectively. The enzyme is composed of six identical subunits. Human TPP‐I has five potential N‐glycosylation sites at Asn residues 210, 222, 286, 313, and 443. A dual role of oligosaccharide at Asn‐286 in folding and lysosomal targeting could contribute to the unusual, but cell type‐dependent, fate of misfolded TPP‐I conformer and represents the molecular basis of the disease process in subjects with naturally occurring missense mutation at Asn‐286 (Wujek et al., 2004). Although TPP‐I zymogen is capable of auto‐activation in vitro, a serine protease that is sensitive to AEBSF participates in the processing of the proenzyme to the mature, active form in vivo (Golabek et al., 2003). TPP‐I is inhibited by PCMBS, DFP, and HgCl2. It is an exo‐type serine peptidase that is regulated by SH reagent. TPP‐I releases the tripeptide Arg‐Val‐Tyr from angiotensin III more rapidly than from Ala‐Ala‐Phe‐ MCA, and also releases Gly‐Asn‐Leu from neuromedin B with the same velocity as from Ala‐Ala‐Phe‐MCA (Du et al., 2001). TPP‐I degrades small peptides with an extended N‐terminal domain, but not structured peptides. In general, this cut off occurs between masses of 4.5 and 6 kDa. Reference to the structures of other peptidases suggests a mechanism for this size selectivity (Bernardini, Warburton, 2001). The order of TPP‐I mRNA expression is as follows: kidney > or ¼ liver > heart > brain > lung > spleen >> skeletal muscle and testis (Du et al., 2001). TPP‐I is largely responsible for the degradation of sulfated cholecystokinin‐8 (CCK‐8S), which enters the cell by receptor‐mediated endocytosis through the cell surface, whereas TPP‐II is responsible for regulating extracellular CCK‐8S levels (Warburton and Bernardini, 2002; Breslin et al., 2003). TPP‐II (EC 3.4.14.10) is a serine peptidase apparently involved in the inactivation of cholecystokinin octapeptide (Rose et al., 1996). TPP‐ II was mostly detected in neurons and also in ependymal cells and choroid plexuses, localizations consistent with a possible participation of the peptidase in the inactivation of cholecystokinin circulating in the CSF. It was also detected at the ultra‐structural level in the cerebral cortex and hypothalamus. The peptidase mainly associated with the cytoplasm of neuronal somata and dendrites, often in the vicinity of reticulum cisternae, Golgi apparatus, or vesicles, and with the inner side of the dendritic plasma membrane (Facchinetti et al., 1999).

6.2 N‐Acetylated a‐Linked Acidic Dipeptidase (NAALADase) NAAG is catabolized to N‐acetylaspartate and glutamate primarily by glutamate carboxypeptidase II (NAALADase), which is expressed on the extracellular surface of astrocytes. NAALADase has been cloned from human brain (Luthi‐Carter et al., 1998a), rat brain cDNA (Luthi‐Carter et al., 1998b), and rat hippocampal cDNA library (Bzdega et al., 1997). NAALADase inhibition prevents cocaine‐kindled seizures (Witkin et al., 2002; Rojas et al., 2003). NAALADase inhibitor 2‐(phosphonomethyl)pentanedioic acid (2‐PMPA) produced dose‐dependent protection (10–100 mg/kg) against both the development of seizure

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kindling and the occurrence of seizures during the kindling process without observable behavioral side effects (Thomas et al., 2001). Its inhibition also protects motor neurons from death in familial amyotrophic lateral sclerosis models (Ghadge et al., 2003).

6.3 Bestatin‐Insensitive Aminopeptidase Bestatin‐insensitive aminopeptidase (BIA), a thiol aminopeptidase, was resistant to puromycin and bestatin inhibition (Neidle et al., unpublished observations). It was purified 744‐fold to homogeneity from rat brain cytosol. BIA was separated from aminopeptidase B (APB) and PSA and isolated by DEAE chromatography. It has a molecular weight of 280 kDa and is composed of six 45‐kDa subunits. It releases Tyr from enkephalin, but not from dipeptides. The enzyme is inhibited by heavy metal ions and thiol‐binding reagents including leupeptin and E‐64. The aminopeptidase can be readily differentiated from enzymes with partially overlapping specificity such as leucine aminopeptidase; aminopeptidases A, B, and N; or PSA by its resistance to inhibition by bestatin and to metal‐chelating agents.

7

Therapeutic Uses of Neuropeptidase Inhibitors

ACE inhibition reduces morbidity and mortality in patients with hypertension, diabetes mellitus, atherosclerosis, heart failure, and nephropathy. ACE inhibitors could be useful in the management of a wide range of cardiovascular pathologies (Stanton, 2003). Patients at risk of cardiovascular events but having normal left ventricular function demonstrate clear benefits of an ACE inhibitor. Patients with chronic left ventricular dysfunction or postmyocardial infarction show reduction of ischemic events (Stanton, 2003). The use of ACE inhibitors to treat hypertension indicates that neuropeptidases do make good drug targets (Docherty et al., 2003). With genome research yielding many possible new drug targets, neuropeptidases that are causally responsible for disease processes might therefore make better targets, especially if it leads to the development of drugs that can be administered orally. Besides inhibitors, antibodies for peptidases can also be useful therapeutics.

7.1 Synthetic Anti‐Enkephalinases Inhibition of aminopeptidases and NEP with bestatin and thiorphan exerts strong and long‐lasting analgesia in terminal cancer patients; a similar effect was found with two other active inhibitors, acetorphan and carbaphethiol (Noble and Roques, 1992). Kelatorphan, a mixed inhibitor of PSA, APN, NEP, and DAP, is as active as morphine in many tests (Thorsett and Wyvratt, 1987). Another mixed inhibitor, phelorphan, affects the morphine withdrawal syndrome (Van Amsterdam et al., 1987). SCH‐34826, an orally active NEP inhibitor that produces analgesia in mice, does not alter gastrointestinal movement or respiratory function (Chipkin et al., 1988). In addition, it is inactive in tests measuring potential antianxiety, antidepression, and antipsychotic effects demonstrating that the inhibitors have more specific pharmacological effects than the opiates. Since the enkephalinergic pathway of analgesia is unique, it is feasible that the NAP inhibitor may be a non‐addictive analgesic without psychotropic side effects.

7.2 Synthetic Anti‐Aminopeptidase The reversible and irreversible inhibitors of aminopeptidases are designed to be small, stable, of higher affinity, and blood–brain barrier permeable (Tieku and Hooper, 1992). The reversible mercaptoethylamine with a hydrophobic side chain (2‐amino‐4‐methyl‐1‐pentanethiol) is a potent dentate inhibitor of APN (Pickering et al., 1985). The reversible carbaphethiol, a parenterally active form of phethiol, was also

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developed. The structural features necessary for interaction of mercaptoamines with aminopeptidases are specific. Aminopeptidases contain subsites that contribute to specific substrate binding: the ‘‘R’’ group of the N‐terminal amino acid and the ‘‘R0 ’’ group of the peptide chain (Bryce and Rabin, 1964; Van Amsterdam et al., 1987). Irreversible inhibitors resistant to metabolic inactivation are of greater value for in vivo studies. Diazomethyl ketone, which specifically attacks the active site cysteine residue (Fujiwara et al., 1982), has been shown to be an extraordinarily potent inhibitor of pyroglutamyl aminopeptidase (Wilk et al., 1985). It is highly effective in in vivo studies and has low toxicity. By substituting the functional groups of these synthetic compounds, potent and selective inhibitors for ENK inactivation may become available.

7.3 Endogenous Peptidase Inhibitors Naturally available inhibitors frequently have higher selectivity and potency than synthetic ones. Snake venom peptide inhibitors played a crucial role in establishing the clinical value of Capoten in hypertension. Microbial inhibitors were shown to be highly active for a spectrum of peptidases and proteinases (Wilk et al., 1985). The discovery of phosphoramidon as a NEP inhibitor has been very helpful in developing novel inhibitors (Kenny, 1977). It is of interest that all the above‐mentioned synthetic and natural inhibitors are modified peptides (Umezawa, 1972; Umezawa and Aoyagi, 1977). Recently one of the most potent ACE inhibitors, designated converstatin, was characterized to be a tryptic fragment of a plasma protein with a Ki value in the picomolar range (Okuda and Arakawa, 1985). Peptide inhibitors for enkephalinases would be superior because of their non‐toxicity (Rapaka, 1986). They can be metabolized to amino acids that do not cause liver and kidney damage as the opiates do. These inhibitors of cerebral peptidases cannot cross the placental barrier, which is an additional advantage for their use as analgesics for pregnant women.

7.4 Endogenous Anti‐Aminopeptidases Arg0‐Met‐Enk was purified and characterized as an endogenous PSA inhibitor in calf striatum (Hui et al., 1994). It is likely that in addition to their possible role as opioids, the enkephalin‐containing polypeptides may be regulators of enkephalin levels (Hui et al., 1982, 1983a). It is the most potent one and is stronger than bestatin and puromycin (Hui et al., 1983b, 1994). The inhibitor can be inactivated by a specific enzyme, DAP‐V, specifically releasing the N‐terminal Arg‐Tyr (Hui, 1988). Although NAP is sensitive to small‐peptide inhibition in vitro, its substrate and inhibitor specificity is different from that of PSA (Hui et al., 1998).

7.5 Inactivation of Arg0‐Met‐Enk Arg0‐Met‐enk can be degraded by a new type of DAP. The enzyme was purified about 2,100‐fold with 7% recovery from the rat brain membrane by column chromatography, successively on Cellux D, Arg‐Tyr‐AH‐ Sepharose 4B, hydroxylapatite, and Sephadex G‐75, after the membrane was solubilized with Nonidet P40 (Hui, 1988). The enzyme activity was assayed by HPLC using Arg0‐Met‐Enk as substrate in the presence of bestatin, thiorphan, and captopril. In SDS‐PAGE, the purified enzyme was apparently homogeneous with a molecular weight of 64,000 daltons. This thiol enzyme is optimally active at pH 7 and is selectively activated by Mnþþ, Coþþ, and Znþþ. It splits Arg0‐Met‐Enk into equal amounts of Arg‐Tyr and Gly‐Gly‐Phe‐Met with a Km of 100 mM and Vmax 3.8 mmol/mg protein per min. DAP does not hydrolyze the model substrates for DAP‐I, DAP‐II, DAP‐III, DAP‐IV, amino acid bNAs, actin, desmin, tubulin, glial fibrillary acidic protein, and cytoskeletonal neurofilament proteins. The enzyme is insensitive to puromycin, but is inhibited by several neruropeptides; angiotensin III is the most potent, with a Ki of 0.3 mM. Its substrate specificity, pH optimum, molecular weight, activators, and catalytic sites demonstrate that this enzyme is distinct from other DAPs.

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Neuropeptidases Regulated by Inhibitors

The presynaptic Enks have to cross the synaptic cleft before acting on the dendrites, axons, or neuronal perikarya (Pasquini et al., 1992). The blood‐borne Enks indicate that some peptide receptors are distant from the synapses. During their travel, attack by peptidases is unavoidable. Recent findings indicate that the pharmacological effects of puromycin, bestatin, and thiorphan are due to their protection of Enk from destruction before its receptor binding and their inhibition of the postsynaptic peptidases (Graf et al., 1982). Bestatin and thiorphan increase the release of Enk by Kþ stimulation (Patey et al., 1981) indicating that peptidase inhibitors play a major regulatory role in the process. The physiological function of insulin is controlled by degradation in its releasing neuron (Halban, 1980). The presynaptic regulation allows the neurons to store releasable peptide that is relatively independent of axoplasmic transport. It is believed that Enks and neurotransmitters alike are constantly released at basal levels, which increase with stimulation. They are continuously produced at a high level and are degraded after production, with degradation being inhibited during stimulation of release. The inhibitor is released together with Enks into the synaptic cleft. Indeed, Enk is co‐released with the peptide (Winkler et al., 1987). A NAP inhibitor is released in rat hippocampal slices by electric stimulation. The enzyme is found to be sensitive to a small‐peptide soluble fraction extracted from bovine brains (Hui and Hui, unpublished observations). The study of the synaptic peptidases and their endogenous inhibitors is important from both a theoretical and practical point of view. First, it enables the design of inhibitors of the relevant peptidase that may mimic, to a large extent, the effects of exogenous opioids. Second, inhibitors may potentiate Enk biological actions, and thereby contribute to the delineation of their functional roles. In addition, it will provide new insight into the regulatory mechanism of peptide neurotransmission in the living brain that will lead to the development of novel inhibitors that are innovative analgesics without the unwanted side effects experienced with narcotics.

9

Conclusion

Neuropeptidases play a critical role in the biosynthesis and metabolism of neuropeptides that are crucial for health. Abnormalities of neuropeptidases have been found in numerous mental, degenerative, and neurological diseases. Opioid peptide enkephalin, the most well studied neuropeptides is synthesized with a group of peptidases acting in an orderly sequence. In contrast, the inactivation of enkephalin is likely to be controlled by a single peptidase—NAP. Study of the neuropeptidases will broaden our knowledge on how the nervous system functions. That will ultimately lead to the development of better diagnosis and treatment, targeted at the gene or enzyme level, in mental and neurological diseases. Therapeutics developed by this approach will be safer, more efficient, and cost effective.

Acknowledgments The work was supported by the Office of Mental Health, New York State, and in part by grants NBS 78‐ 26164 from the National Science Foundation, and NIDA 5R01DA06271 and NIMH S15MH51893 from the National Institutes of Health.

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Ubiquitination and Proteasomal Protein Degradation in Neurons

L. Klimaschewski

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 654 2 The Ubiquitin‐Proteasome System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 654 3 Ubiquitination and Deubiquitination in Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 655 4 Subunit Composition and Regulatory Functions of the Neuronal Proteasome . . . . . . . . . . . . . . . . 657 5 The Crucial Role of the UPS in Neurological Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 658 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 659

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Ubiquitination and proteasomal protein degradation in neurons

Abstract: The ubiquitin‐proteasome system (UPS) is responsible for regulated protein degradation in all eukaryotic cells. In nerve cells, this pathway determines protein abundance and protein quality control in spatially restricted neuronal compartments, such as the nucleus, dendrites and spines, axons, and in synaptic boutons. Therefore, the UPS regulates neuronal gene expression, synaptic and spine functions, axonal regeneration, and neuronal degeneration by tagging and eliminating key proteins required for morphological and chemical neuroplasticity. Recent progress in the elucidation of the pathomechanisms leading to various neurological disorders clearly revealed the importance of the UPS for neuron survival and maintenance. The significance of ubiquitin‐mediated proteolysis in neuronal physiology and pathophysiology has now become firmly established and brings this pathway into the focus of many neuroscience laboratories worldwide. List of Abbreviations: DUB, deubiquitinating enzyme; E1, ubiquitin-activating enzyme; E2, ubiquitinconjugating enzyme or ubiquitin carrier; E3, ubiquitin ligase; ROS, reactive oxygen species; UBL, ubiquitinlike protein; UBP, ubiquitin-specific processing enzyme; UCH, ubiquitin carboxy-terminal hydrolase; UCH, ubiquitin C-terminal hydrolase; UPS, ubiquitin-proteasome system

1 Introduction Protein modification by the 76‐amino‐acid residue ubiquitin (ubiquitination) or by one of the ubiquitin‐ like (UBL) proteins (e.g., SUMO or Nedd8) reflects a cellular mechanism of posttranslational protein regulation similar to protein phosphorylation. Ubiquitination functions as a targeting signal making it possible for the proteasome to recognize and destroy the protein tagged with ubiquitin. In addition to eliminating abnormally folded or mutated proteins, ubiquitination targets ER lumenal and membrane proteins for degradation in the lysosome (for reviews see Jentsch (1992); Hochstrasser (1995); Hershko and Ciechanover (1998); Hicke (2001); Welchman et al. (2005)).

2 The Ubiquitin‐Proteasome System Multiubiquitination (several ubiquitins attached to each other) marks proteins for degradation in the proteasome. In contrast, monoubiquitination (attachment of a single ubiquitin) generally indicates a conformational change in the protein, while polyubiquitination (multiple monoubiquitination) labels plasma membrane proteins for endocytosis. Ubiquitination is accomplished by a series of three enzymatic steps (> Figure 22‐1). These involve the ATP‐dependent activation of ubiquitin by the ubiquitin‐activating enzyme (E1), transfer of ubiquitin to one of the 25–30 ubiquitin‐conjugating enzymes (E2 or ubiquitin carriers), and isopeptide linkage of ubiquitin’s C‐terminal glycine to the e‐amino group of lysine residues within the target protein. This last step is catalyzed by an ubiquitin ligase (E3). Differences among E2s, which are characterized by a 14–16 kDa conserved core domain, determine the specificity of their interactions with one of the approximately 1000 E3s (i.e., the number of possible E2/E3 combinations is in the same range as the approximately 30,000 gene products encoded in the human genome). Some E2s have amino‐ or carboxy‐terminal extensions that facilitate interactions with ubiquitin ligases. The latter confer specificity in the ubiquitination process. They are characterized by different subdomains (‘‘HECT’’ or ‘‘RING’’). HECT domains represent the 350‐amino‐acid residue carboxy‐ terminal sequence exhibiting a conserved cysteine for intermediate binding of ubiquitin, whereas RING fingers consist of eight metal‐binding residues coordinating two zinc ions and serve as the docking site for the ubiquitin‐conjugating enzyme (E2). Finally, complex multisubunit E3s have been described. Besides detecting specific signals in the target protein, ubiquitin ligases bind to chaperones to recognize misfolded proteins. The E3 CHIP, for example, ubiquitinates hyperphosphorylated tau protein in axons via interaction with the Hsc70 chaperone (Shimura et al., 2004). Multiubiquitin chains of four or more ubiquitin molecules (linked through lysine 48) are recognized by the 26S proteasome, a large cylindric complex consisting of more than 60 subunits. The recognition of the


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. Figure 22‐1 Protein ubiquitination and degradation by the ubiquitin‐proteasome system (UPS) and its possible role in neuronal toxicity. See text for abbreviations. Adapted from Klimaschewski (2003)

ubiquitinated target protein is mediated by the regulatory 19S particle containing ATPases, ubiquitin‐ binding subunits, and a deubiquitinating enzyme (DUB). The 20S moiety, in contrast, is composed of four stacked rings (each with seven subunits). The multiubiquitinated protein is unfolded and cleaved by proteolytic scissors at the C‐terminal end of large hydrophobic, basic, or acidic amino acids. The three catalytic activities are buried in the inner rings of the 20S core. The degradation products—short peptide fragments and amino acids—are reused for protein synthesis, while multiubiquitin chains are disassembled through the activity of DUBs that act as cysteine proteases like the ubiquitin carboxy‐terminal hydrolases (UCHs) or ubiquitin‐specific processing enzymes (UBPs). Deubiquitination has emerged as a tightly regulated process, unlike the removal of phosphate groups by phosphatases. Recent evidence points to an important function of ubiquitin C‐terminal hydrolase as an immediate early gene involved in long‐term facilitation in Aplysia neurons.

3 Ubiquitination and Deubiquitination in Neurons Ubiquitin is highly expressed during neuronal development in dendrites of early postnatal pyramidal neurons in the cortex and hippocampus as well as in Purkinje cells of the cerebellum (Flann et al., 1997). However, the distribution and functional significance of ubiquitin carriers, ubiquitin ligases, or DUBs in subcellular neuronal compartments has not been elucidated until recently (for review see Klimaschewski (2003)). Neuroscientists for many years applied antibodies against the neuronal marker protein, PGP9.5, to label central and peripheral neuronal cell bodies and their axons. This antigen was identified as a highly conserved ubiquitin C‐terminal hydrolase (UCH) with specific expression in neurons and was estimated to comprise 1–2% of the total soluble brain protein. In Aplysia, learning induces ap‐UCH that associates with the proteasome and increases its proteolytic activity (Hegde et al., 1997). Inhibiting the hydrolase blocks induction of long‐term but not short‐term facilitation. This may be explained by increased degradation of inhibitory constraints on long‐term memory storage. In general, however, increases in activities of E3s or of the proteasome are associated

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with synaptic degradation (pruning), while synapse strengthening is accompanied by blockade of ubiquitin ligation pathways and reduced proteasomal activity resulting in the stabilization of substrates required for synapse growth. Synaptic stimulation induces rapid changes in the ubiquitination of proteins within the growth cone (Chen et al., 2003). This process has been further investigated in Drosophila suggesting that synaptic growth at the neuromuscular junction is reciprocally regulated by the DUB Fat facets (faf) that increase the number of synpatic boutons (DiAntonio et al., 2001), and by the E3 Highwire (hiw) that destabilizes synaptic contacts (Wan et al., 2000). Spine stability is regulated by the UPS as well. This process involves proteasomal degradation of SPAR (spine‐associated Rap GTPase activating protein) leading to spine loss via decreased levels of postsynaptic density proteins like PSD‐95 (Pak and Sheng, 2003). During neuronal differentiation in vitro, the pheochromocytoma PC12 cell line (Greene and Tischler, 1982) exhibits increased levels of high‐molecular weight ubiquitin‐conjugates in PC12 nuclei, decreased levels of free ubiquitin in cytoplasmic and nuclear extracts, upregulation of at least four E2 activities, and enhanced capacities for ubiquitination (Takada et al., 1994; Ohtani‐Kaneko et al., 1996; Obin et al., 1999). In contrast, proteasomal activity is downregulated when compared with proliferating PC12 cells (Klimaschewski et al., 2006). These findings suggest that during neurite outgrowth nuclear proteins are ubiquitinated at an enhanced rate and, thereby, targeted for degradation. The search for enzymes responsible for increased utilization of ubiquitin revealed the ubiquitin‐ conjugating enzyme, HR6B (yeast UBC2/RAD6), whose mRNA and protein levels increase in PC12 cells after treatment with nerve growth factor (NGF) (Kavakebi et al., 2005). HR6B is highly homologous to the bendless gene (ben), which was originally isolated as a fruit fly missense mutation affecting the escape jump response of Drosophila (Muralidhar and Thomas, 1993). Ben is expressed during development and if mutated leads to defects in synaptic connectivity particularly within the giant fiber circuitry (Oh et al., 1994). HR6B participates in ‘‘N‐end rule degradation’’ implicated in the cleavage of proteins with destabilizing N‐terminal residues (bulky hydrophobic or basic amino acids) and requires UBR1, the ubiquitin ligase that binds N‐end rule target proteins. Downregulation of HR6B or UBR1 mRNAs inhibits neurite outgrowth by PC12 cells. Furthermore, axonal regeneration in cultured primary neurons, which express prominent nuclear and membrane‐associated HR6 immunoreactivity, is reduced by dipeptide‐inhibitors of UBR1 in vitro (Kavakebi et al., 2005). Therefore, N‐end rule ubiquitination appears to be required for neuronal differentiation and may be involved in the axonal regeneration of peripheral neurons. In fact, ubiquitin mRNA is elevated after axotomy, suggesting enhanced requirement for ubiquitin during axonal regeneration (Savedia and Kiernan, 1994). In addition to UBR1, other ubiquitin ligases have been demonstrated to be responsible for the degradation of proteins involved in axon growth. For example, endogenous downregulation of RhoA, a small GTPase highly relevant for dendritic growth and axonal regeneration in central and peripheral neurons (Kaibuchi et al., 1999), is mediated by the ubiquitin ligase, Smurf1 (Wang et al., 2003). Smurf1 enhances neurite outgrowth in Neuro2a neuroblastoma cells, and Smurf1 overexpression reduces RhoA protein levels during cAMP‐induced neurite outgrowth (Bryan et al., 2005). Therefore, decreasing RhoA levels via enhancing its ubiquitination represents a novel way to promote neurite outgrowth. Vice versa, other targets stimulating axon growth are probably ubiquitinated by the anaphase‐promoting complex (APC), an E3 acting as a regulator of mitosis, which is detected in the nuclei of postmitotic neurons in all brain regions indicating important functions beyond the control of cell‐cycle progression (Gieffers et al., 1999). Inhibition of Cdh1‐APC in primary neurons specifically enhances neurite outgrowth and overrides the inhibitory influence of myelin on axon regeneration (Konishi et al., 2004). Another E3 with relevance for neuronal development and maintenance, the RING finger ubiquitin ligase c‐Cbl, is required for ubiquitination of receptor tyrosine kinases and of other membrane proteins like E‐cadherin. Receptor tyrosine kinases mediate the effects of various neurotrophic factors on survival and process growth by peripheral and central neurons (Dechant et al., 1994). In response to ligand binding, dimerization, and activation, tyrosine kinase receptors initiate a cascade of events that include negative signaling aimed at decreasing the amplitude of positive signals, and thus modulate the level of growth factor stimulation. Activation induces receptor monoubiquitination at multiple sites by c‐Cbl, which targets the receptor for endocytosis via clathrin‐coated pits. While still active as part of the endosomal membrane, the receptor is then sequestered into intraluminal vesicles of multivesicular bodies and subsequently degraded


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by lysosomal enzymes (Bache et al., 2004). In the case of fibroblast growth factor signaling, however, c‐Cbl does not directly bind to the receptor but catalyzes the ubiquitination of the receptor via interaction with FRS2 and Grb‐2 (Wong et al., 2002). Increasing evidence suggests that ubiquitination of membrane receptors (‘‘regulated endocytosis’’) targets them for degradation, whereas ubiquitin‐independent internalization (‘‘constitutive endocytosis’’) results in recycling back to the plasma membrane. Interestingly, although the proteasome is not expected to participate in the degradation of membrane proteins directly, inhibition of the proteasome prevents endocytosis and degradation of various neuronal receptors including opioid‐, vasopressin‐, AMPA‐ and GABA receptors (reviewed in Hegde (2004)). In summary, monoubiquitination has now been clearly established as a frequently used ‘‘trafficking signal’’ for membrane‐ associated proteins that shuttle between different vesicular compartments before they are finally degraded by the multivesicular/lysosomal pathway in a proteasome‐dependent manner.

4 Subunit Composition and Regulatory Functions of the Neuronal Proteasome Proteasomal subunits are differentially expressed between organs, i.e., proteasomes from the brain are different from those found in the spleen (Noda et al., 2000). Immunoreactivity against the 20S proteasome particle is heterogenously located in all brain areas with predominant staining of large motor neurons (Mengual et al., 1996). All neuronal compartments contain proteasomes, indicating a functional role of localized protein degradation in various aspects of neuronal morphology (Guo et al., 2001). Cytosolic proteasomal activity accounts for most of the total proteasomal activity, although nuclei exhibit strong immunoreactivity for proteasomal subunits in the brain. Proteasomal subunits with catalytic activities are regulated upon neuronal differentiation in human neuronal progenitor cells as well as in microglial cells by interferon and other inflammatory agents resembling the exchange of subunits in the antigen‐presenting cells of the immune system (Stohwasser et al., 2000). Therefore, proteasomal subunit composition is modulated in response to the specific needs of neurons and glial cells. Regulating protein levels via modification of proteasomal activity may play an important role in neuronal development. Inhibition of the proteasome by lactacystin, a specific blocker of the catalytic b‐ subunits, results in transient neurite outgrowth by neuronal cell lines, an effect probably mediated by stress‐ activated protein kinases (Giasson et al., 1999). Inhibition of other proteases like calpain does not promote neuronal differentiation, indicating the specificity of the lactacystin response (Ohtani‐Kaneko et al., 1998). Interestingly, treatment of the pheochromocytoma PC12 cell line with NGF or with other differentiating agents reduces proteasomal activity. This was shown to be accompanied by an increase in mRNA and protein levels of the catalytically active subunits b1, b2, and b5, but not of their inducible counterparts (Klimaschewski et al., 2006). Moreover, direct phosphorylation of the proteasomal b6‐subunit by the NGF receptor tyrosine kinase A (trkA) was demonstrated (MacDonald et al., 1999). These findings clearly indicate changes in the subunit composition and the activation state of the proteasome during neuronal development and differentiation. In contrast to neuronal cell lines, however, pretreatment of primary neurons with proteasome inhibitors completely prevents axon formation, and lower concentrations of lactacystin (0.5–5 mM) significantly reduce axonal elongation and branching in vitro (Klimaschewski et al., 2006). Established axonal networks degenerate rapidly, and long‐term survival of peripheral neurons is impaired in the presence of proteasome inhibitors. Axonal pathology is reminiscent of the morphological changes in neurodegenerative disorders and supports a crucial role of the three constitutively active catalytic subunits in axon initiation, maintenance, and regeneration. The important role of the proteasome in axon guidance and growth cone dynamics is further corroborated by several studies investigating guidance molecules like netrins, semaphorins, and ephrins, for example, proteasome inhibitors block netrin‐1‐induced chemotropic responses (for review see Patrick (2006)). Another example is provided by the midline repellent Slit and its receptor Robo (Roundabout). Downregulation of Robo is mediated by ubiquitination of the adaptor protein Commissureless (Comm) via the ubiquitin ligase Nedd4, allowing axons to cross the midline (Myat et al., 2002).

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Complete and irreversible blockade of the proteasome by lactacystin ultimately arrests cellular metabolism and causes cell death. In fact, various indicators of neuronal apoptosis including chromatin fragmentation, DNA laddering, caspase activation, poly(ADP‐ribosylation), and release of cytochrome c have been observed in response to proteasome inhibition (for review see Ding and Keller (2001)). However, depending on the cell type and concentration of the respective proteasomal inhibitor, the onset of apoptosis is delayed in some neuronal in vitro models, probably because proteasomal activity is required for the activation of proapoptotic signaling intermediates (Maggirwar et al., 1998). It has been shown that lactacystin blocks caspase‐1‐mediated processing of poly(ADP‐ribose) polymerase (PARP), thus prolonging survival of postmitotic NGF‐deprived sympathetic neurons (Sadoul et al., 1996). Furthermore, proteasome inhibitors prevent early caspase‐3 activation and tau cleavage in cerebellar granule neurons undergoing cell death induced by reduced extracellular potassium (Canu et al., 2000). In this model, however, UPS function is significantly impaired in the later execution phase of neuronal apoptosis, accompanied by an accumulation of ubiquitinated and aggregated proteins that may physically obstruct proteasome entry or block the catalytic sites within the inner rings of the central core. Inhibition of the proteasome may result from reactive oxygen species (ROS) as well. ROS have been suggested to play an important role in proteasome inhibition by increasing the level of lipid peroxidation products followed by protein cross‐linking. This may prevent unfolding of peptides before they are transferred into the catalytic chamber of the proteasome, thereby effectively acting as dominant inhibitors. ROS, however, serve a number of essential intracellular functions as signaling intermediates and may even enhance proteasomal activity under some conditions (Reinheckel et al., 1998). Moreover, neurons may activate protective mechanisms against high concentrations of ROS by increasing levels of heat shock proteins, which aid in folding, trafficking, or disaggregation of proteins on their way to the proteasome. Furthermore, the proteasome can be reversibly blocked for short periods of time by physiological inhibitors like PI31 to promote degradation of oxidized proteins in the lysosomal pathway instead (Zaiss et al., 1999).

5 The Crucial Role of the UPS in Neurological Disease Some ubiquitin ligases have been identified in their mutated form causing rare, inheritable neurological diseases. Angelman’s syndrome is a congenital disorder caused by mutations in the E6‐AP gene that encodes a HECT ubiquitin ligase (Kishino et al., 1997). Affected children exhibit movement disturbances, inability to speak, and mental retardation. More common neurological disorders like Parkinson’s disease are also associated with mutations in components of the UPS. Autosomal recessive forms of Parkinson’s disease (arPD) are at least in part caused by mutations in parkin, a member of the RING finger family E3s (Shimura et al., 2000). Substrates of parkin have been identified, such as the cell division control protein (CDCrel‐1), a member of the septin GTPase family. Furthermore, the parkin‐associated endothelin‐receptor‐like receptor (Pael‐R) induces endoplasmic reticulum stress and elicits cell death when overexpressed, whereas coexpression of intact parkin protects against this death (Imai et al., 2001). Other substrates of parkin probably include the glycosylated form of a‐synuclein (Sp22) and synphilin‐1 (for review see Ross and Pickart (2004)). It has to be pointed out, however, that mice with targeted deletion of parkin do not exhibit elevated levels of the candidate target proteins, suggesting problems with the overexpression approach (Goldberg et al., 2003). Moreover, a‐synuclein may directly be recognized by the proteasome without prior tagging by ubiquitin. In addition to parkin, the ubiquitin hydrolase UCH‐L1 and DJ‐1—a substrate for ligation to the ubiquitin‐related modifier SUMO—have been linked to rare forms of Parkinson’s disease. DJ‐1 probably acts as a redox‐sensitive chaperone that becomes unstable and degraded in its mutated form, while the Ile93Met mutation in UCH‐L1 results in Parkinson’s disease by decreasing the enzyme’s activity (Leroy et al., 1998). As discussed below, it appears likely that failure in ubiquitination and deubiquitination as well as general proteasome inhibition make catecholaminergic neurons particularly vulnerable to cellular stress and disturbances in the balance of the redox system (Dawson and Dawson, 2003). Neurodegenerative disorders like Parkinson’s or Alzheimer’s disease, which account for the cognitive decline of the majority of elderly patients, are characterized by the accumulation of intraneuronal


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inclusions, which may be regarded as neuropathological hallmarks of other human chronic neurodegenerative disorders as well (Alves‐Rodrigues et al., 1998). The proteinaceous, insoluble aggregates are detected in the neuronal cytoplasm, axoplasm, and in neuronal nuclei. They may be encapsulated by intermediate filaments. Recent evidence suggests that inclusions are formed through cross‐linking of various hydrophobic regions exposed by several mechanisms, e.g., by mutation, misfolding, or oxidative damage, and retrogradely transported along microtubules (Kopito, 2000). Examples for cytoplasmic inclusions represent neurofibrillary tangles in Alzheimer’s disease or Lewy bodies in Parkinson’s disease. Nuclear inclusions are observed in expanded polyglutamine disorders, for example, in spinocerebellar ataxia type 1 (SCA1). SCA1 is caused by mutant ataxin‐1, which is ubiquitinated like normal ataxin‐1 but resists proteasomal degradation. The HECT ubiquitin ligase E6‐AP is apparently necessary for inclusion formation in SCA1, because mice exhibiting expanded polyglutamine ataxin‐1 and simultaneously lacking E6‐AP reveal significantly fewer nuclear inclusions (Cummings et al., 1999). The role of the UPS in inclusion body formation became clear when a variety of molecules involved in ubiquitin‐mediated protein degradation, among them proteasomal subunits and several components of the UPS, were demonstrated in these aggregates. Ubiquitin‐positive inclusions are detected within the neuronal nucleus of patients with Huntington’s disease (Sieradzan et al., 1999) suggesting that abnormal huntingtin protein (containing expanding CAG repeats) is targeted for proteolysis but resistant to effective removal (Waelter et al., 2001). Increasing evidence now suggests that proteins forming aggregates either resist or inhibit proteasome‐mediated proteolysis (> Figure 22‐1), and mutations that impair normal ubiquitination (or deubiquitination) may lead to protein accumulation resulting in proteasomal inhibition (Bence et al., 2001). Vice versa, inhibition of the proteasome by blockade of the catalytic activities results in inclusion formation (Rideout et al., 2001). Further support for this hypothesis is provided by studies demonstrating that mutations in ubiquitin ligases such as parkin reduce the formation of ubiquitin‐positive inclusions. Therefore, segregation of abnormal proteins requires ubiquitination and serves as a natural defense mechanism against the cytotoxic effects of abnormal proteins that cannot be recognized and degraded by the proteasome (McNaught et al., 2001). Lewy bodies frequently occur in late‐onset sporadic Parkinson’s disease, while they are rarely found in patients with young‐onset familial forms of Parkinson’s that is often accompanied by severe neurodegeneration. The high number of cases with incidental Lewy bodies and mild neurodegeneration in the elderly probably represents a preclinical stage of Parkinson’s disease characterized by a still intact ability of the neuron to compartmentalize abnormal or cytotoxic proteins as insoluble aggregates in inclusions. From studies in postmortem extracts of the brain using fluorogenic peptide substrates it became clear that proteasomal activity is inhibited under pathological conditions like Parkinson’s, Lewy body formation, or Alzheimer’s disease as well as in ischemia–reperfusion injury or during aging (Ding and Keller, 2001). Furthermore, prolonged inhibition of proteasomal activity causes accumulation of various proteins including cyclin D1, a marker of the G1/S transition of the cell cycle, which induces cell death when overexpressed in neuronal lines (Boutillier et al., 1999). Moreover, disease‐associated proteins such as presenilin, hyperphosphorylated neurofilaments, amyloid peptide, and parkin are elevated if proteasomes are blocked. The vicious cycle probably starts when accumulated proteins block the internal compartment of the proteasome, thereby further reducing proteasomal activity and initiating the cell death program. On the other hand, the pathogenetic mechanism of neuronal apoptosis in neurodegenerative diseases may be far more complex, since blockade of ubiquitination or of the proteasome may prevent the neurotoxicity of accumulated proteins like the amyloid peptide as well (Favit et al., 2000).

6 Conclusions Dissecting the function of ubiquitination, deubiquitination, and proteasomal degradation in neuronal differentiation, apoptosis, and disease provided the basis for our current understanding of the UPS in the physiology and pathology of the nervous system. The mechanisms involved in the regulation of the half‐life of proteins required for process outgrowth, synaptic plasticity, neuronal survival, and overall maintenance of neural cells are now at the core of neuroscience research laboratories around the world. From a clinical

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point of view, the UPS clearly plays a key role in the pathogenesis of several—if not all—neurodegenerative disorders. Some mutated or misfolded proteins probably resist ubiquitination or recognition of their ubiquitinated forms by the proteasome. It is becoming evident that segregation of these proteins into inclusion bodies evolved as a survival mechanism to prevent the toxicity of these proteins in their soluble or protofibrillary form.

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mouse microglia: Induction of immunoproteasomes by interferon‐g and lipopolysaccharide. Glia 29: 355-365. Takada K, Kanda T, Ohkawa K, Matsuda M. 1994. Ubiquitin and ubiquitin‐protein conjugates in PC12h cells: Changes during neuronal differentiation. Neurochem Res 19: 391398. Waelter S, Boeddrich A, Lurz R, Scherzinger E, Lueder G, et al. 2001. Accumulation of mutant huntingtin fragments in aggresome‐like inclusion bodies as a result of insufficient protein degradation. Mol Biol Cell 12: 1393-1407. Wan HI, Di Antonio A, Fetter RD, Bergstrom K, Strauss R, et al. 2000. Highwire regulates synaptic growth in Drosophila. Neuron 26: 313-329.

Wang HR, Zhang Y, Ozdamar B, Ogunjimi AA, Alexandrova E, et al. 2003. Regulation of cell polarity and protrusion formation by targeting RhoA for degradation. Science 302: 1775-1779. Welchman RL, Gordon C, Mayer RJ. 2005. Ubiquitin and ubiquitin‐like proteins as multifunctional signals. Nat Rev Mol Cell Biol 6: 599-609. Wong A, Lamothe B, Lee A, Schlessinger J, Lax I, et al. 2002. FRS2 a attenuates FGF receptor signaling by Grb2‐ mediated recruitment of the ubiquitin ligase Cbl. Proc Natl Acad Sci USA 99: 6684-6689. Zaiss DM, Standera S, Holzhutter H, Kloetzel P, Sijts AJ. 1999. The proteasome inhibitor PI31 competes with PA28 for binding to 20S proteasomes. FEBS Lett 457: 333-338.

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Protease Activity in the Aging Brain

D. A. Gray

1 Proteolytic Systems in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 664 2 Age‐Related Changes in the Lysosomal System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 665 3 Age‐Related Changes in the Ubiquitin‐Proteasome System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 665 4 The Garbage Catastrophe Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 666 5 Genetic Models of Age‐Related Proteolytic Deficit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 667 6 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 669

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Protease activity in the aging brain

Abstract: There is abundant evidence for the accumulation of damaged and misfolded proteins in the aging mammalian brain. This accumulation may result from an increasing burden of oxidative damage combined with a diminished capacity to degrade aberrant proteins through the lysosomal and proteasomal pathways (the two major systems for the regulated disassembly of proteins). A chronic proteolytic deficit in either or both systems is predicted to lead to a neurotoxic crisis that has been described as ‘‘garbage catastrophe.’’ The evidence for the decline in proteolytic capacity of the degradative systems and the hypothesized events leading up to their catastrophic failure are reviewed. List of Abbreviations: Atg, autophagy related gene; ATPase, enzyme catalyzing cleavage of adenosine triphosphate; ax, ataxia; CA, Cornu Ammonis (Ammon’s horn), a region of the hippocampus; CLN, ceroid lipofuscinosis, neuronal; CNS, central nervous system; DG, dentate gyrus, a region of the hippocampus; ERAD, endoplasmic reticulum associated degradation system; H, Hilus, a region of the hippocampus; Hsc, cytosolic heat shock protein; KFERQ, motif targeting proteins to the lysosome (refers to single letter amino acid code); NCL, neuronal ceroid lipofuscinosis; PPT, palmitoyl protein thioesterase; ROS, reactive oxygen species; UCH, ubiquitin carboxyterminal hydrolase; UPS, ubiquitin proteasome system; Usp, ubiquitin specific protease; 19S, cap structure of the proteasome (19S refers to centrifugal sedimentation value); 20S, catalytic core of the proteasome (20S refers to centrifugal sedimentation value); 26S, complete proteasome assembly with core rings and end caps (26S refers to centrifugal sedimentation value)

1 Proteolytic Systems in the Brain Cells in the brain are endowed with a large repertoire of proteases, but the current discussion concentrates only on those dedicated to the complete degradation of protein substrates, as opposed to those involved in incomplete proteolysis of substrates through cleavage at specific recognition sites (the role of proprotein convertases, caspases, and so forth). A proximal (and essential) goal of proteolysis may be the timely removal of damaged proteins or those whose degradation is necessary for the cell cycle, signaling cascades, and so forth, but the ultimate (and equally important) goal of the systems under consideration here is the recovery of molecules that can be reutilized in protein synthesis. For this purpose, the cell has two major systems, one compartmentalized within membrane‐bound structures in the cytoplasm, and the other distributed throughout the cytosol and nucleus. The proteases of the former system are concentrated within lysosomes, organelles bound by a single membrane to which substrates are directed by endosomes or phagosomes derived from the plasma membrane. The special case of autophagosomes (double‐membraned structures in which cell constituents are engulfed and delivered to the lysosome for disassembly and recycling) has particular relevance to aging and is considered in a subsequent discussion of the ‘‘garbage catastrophe.’’ The lysosomal system is responsive to environmental conditions, reducing the half‐life of selected substrates when nutrients are limiting (Dice et al., 1986). The lysosomal system demonstrates a degree of substrate specificity; proteins with the KFERQ motif are targeted to the lysosome and hence are favored substrates under stress conditions (reviewed in Dice et al. (1990)). Lysosomal degradation of whole organelles cashiered from service by autophagy may represent the only means by which the cell can reclaim vital resources from complex macromolecular assemblies for reutilization elsewhere. In situations where the organelle has been damaged and may present abnormal molecular structures the broad spectrum of protease activity within the lysosome would be advantageous. The second major proteolytic system within cells is the ubiquitin‐proteasome system (UPS). As in the lysosomal system, the proteases of the UPS have the capacity to degrade most protein substrates, but whereas the proteases of the former are confined to membrane‐bound vesicles the enzyme active sites of the latter are sequestered to the interiors of 20S proteasome core units, cylindrical stacks of rings at the center of the 26S proteasome (reviewed in Wolf and Hilt (2004)). The ports through which substrates access these chambers have a diameter sufficient to accommodate the linear polypeptide chain, but proteins with stable tertiary structure cannot enter this chamber in their folded state. Such proteins must first be unfolded by the action of ATPase enzymes resident in the 19S cap structures of the 26S proteasome. The degradation of


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undamaged, folded protein substrates by the proteasome is brought about by the assembly on these substrates of ubiquitin chains of particular topology; such chains have physical affinity for the proteasome and will deliver the substrate for degradation (such would be the case for proteins whose degradation is required for cell‐cycle progression, the propagation of signaling cascades, the regulation of transcription, and so forth). Proteins that inherently have little tertiary structure or that are damaged by oxidation may be degraded by the proteasome in the absence of ubiquitin‐mediated targeting; in vitro such substrates can be degraded by the 20S core (without the unfolding ATPase activity of the 19S end caps) (Grune et al., 1997; Sheaff et al., 2000; Tofaris et al., 2001).

2 Age‐Related Changes in the Lysosomal System Declining efficiency of lysosome‐mediated proteolysis may arise either as a consequence of inefficient delivery of substrates to the lysosome or as a loss of proteolytic capacity within the organelle itself. There is evidence that the endosomal uptake of plasma membrane proteins is less efficient in aging neurons compared with their younger counterparts (Blanpied et al., 2003), not as a consequence of reduced abundance of endosomal components but rather owing to reduced rates of transport. The delivery of secretory vesicles to the lysosome rather than to the plasma membrane (a mechanistically distinct process designated crinophagy) has not been extensively studied in the context of aging, and it is not yet known to what extent this process is affected in the aging neuron. The delivery of proteins containing the KFERQ motif to the lysosome occurs via direct transport through the lysosomal membrane. This process is mediated by a cytosolic chaperone protein (hsc73) which together with the substrate binds a membrane receptor (lamp2a); on the matrix side of the membrane the process is assisted by a second chaperone, ly‐hsc73 (Cuervo et al., 1997). The activity of chaperone‐mediated lysosomal degradation is regulated by the availability of lamp2a, which is itself tightly regulated by lysosomal degradation (Cuervo and Dice, 2000a). The reduction in chaperone‐mediated degradation observed in senescent fibroblasts (Dice, 1982) may be a consequence of reduced levels of lamp2a (discussed in Cuervo and Dice (2000b)); it remains to be seen whether this process is altered in the aging brain. Macroautophagy involves the engulfment of complete organelles (mitochondria, peroxisomes) by a double‐membraned structure (the autophagic vacuole), which acidifies and fuses to the lysosome. In general terms, it is a mechanism whereby cell constituents can be salvaged and redeployed in times of starvation. The efficiency of macroautophagy declines with age as fewer autophagosomes are formed and the rate of fusion with lysosomes is diminished (Terman, 1995). Within the lysosome, proteolytic cleavage of substrates is mediated by enzymes of the cathepsin family of proteases. The age‐related changes in cathepsin abundance within the CNS have been investigated by Nakanishi and coworkers and have been summarized in an admirable review (Nakanishi, 2003). Whereas the levels of cathepsins D and E were found to increase in the aged rat brain, the enzymatic activity of cathepsin B was increased in the neostriatum but unchanged in other brain regions (Nakanishi et al., 1994, 1997). In contrast, the enzymatic activity of cathepsin L was significantly reduced (approximately 90%) in all regions of the aging rat brain (Nakanishi et al., 1994). The imbalance of cathepsin activities during aging and age‐related diseases is hypothesized to provoke deleterious effects on CNS neurons (Nakanishi, 2003).

3 Age‐Related Changes in the Ubiquitin‐Proteasome System The UPS is charged with the rapid degradation of substrates including oxidized proteins, which are aggregation prone and potentially toxic. Degradation of proteins via the UPS requires the orchestration of complex enzymatic events (reviewed in Passmore and Barford (2004)), some of which may decline in efficiency with age. If the erosion of proteasome activity contributes to aging (as opposed to being merely symptomatic of aging) the following three predictions can be made: (1) There should be an age‐related decrease in proteasome activity and/or proteasome abundance; (2) Artificial inhibition of the proteasome should accelerate aging in whole or in part; (3) Enhancement of proteasome activity should forestall aging

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in whole or in part. Several laboratories have studied alterations in proteasome activity with aging and have documented a decline in tissues such as heart (reviewed in Bulteau et al. (2002)) and skeletal muscle (Husom et al., 2004), skin (Bulteau et al., 2000; Petropoulos et al., 2000), lens (Viteri et al., 2004), and the brain (reviewed in Gray et al. (2003)) as well as in immune cells such as lymphocytes (Ponnappan et al., 1999; Carrard et al., 2003). The decrease in enzymatic activity is typically in the order of 1.5‐ to 2‐fold, and there is evidence correlating this decline with loss of specific proteasome subunits. Studies from the laboratory of Gonos indicate that whereas the a subunits of the proteasome (noncatalytic subunits that assemble into heptameric rings and scaffold the subsequent assembly steps) are in excess and do not decline with age, the catalytic subunits of heptameric b rings do demonstrate an age‐related decline in abundance, and may limit the proteolytic capacity of aging cells (Chondrogianni and Gonos, 2005). Intriguingly, the proteasomes of centenarians (individuals in whom robust systems for the avoidance or repair of molecular damage can reasonably be presupposed) were found to have proteolytic systems commensurate with lesser age (Chondrogianni et al., 2000). In cultures of primary cells, proteasome inhibition was found to induce cellular senescence (Chondrogianni and Gonos, 2004) whereas overexpression of the b5 subunit increased resistance to oxidative stress and increased replicative potential by several population doublings (Chondrogianni et al., 2005).

4 The Garbage Catastrophe Model The term ‘‘Garbage Catastrophe’’ was coined by Terman (2001) to describe the situation that arises when proteolytic systems are unable to eliminate waste toxic to those very systems; a feedback inhibitory loop is generated wherein the accumulating waste promotes its further accumulation, leading to catastrophic failure of proteolysis. The problem is postulated to be a particular threat to postmitotic cells such as cardiac myocytes or neurons (without the potential to grow and divide such cells cannot reduce the burden of nondegradable waste through simple dilution). The waste product implicated in the garbage catastrophe hypothesis is lipofuscin (‘‘aging pigment’’), a complex of cross‐linked proteins, lipids, and heavy metals that may in large part be derived from oxidatively damaged mitochondria (von Zglinicki et al., 1995; Terman and Brunk, 1998b; Brunk and Terman, 2002; Terman and Brunk, 2004) (> Figure 23-1). It is possible to generate a substance with properties very much like lipofuscin (in terms of composition, autofluorescence, etc.) in the laboratory simply by extensively cross‐linking mitochondria with ultraviolet . Figure 23-1 Anterior horn motor neurons in the spinal cord of a 73‐year‐old male with no history of neurological illness. The section was stained with hematoxylin and eosin as well as with periodic acid–Schiff. Note the accumulations of numerous lipofuscin granules (arrows) in the cytoplasm of the neurons


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irradiation (von Zglinicki et al., 1995; Gray and Woulfe, 2005). It is unlikely that the damaged mitochondrial components within lipofuscin would be competent for electron transport etc., but within the environment of the lysosome, the heavy metals trapped therein could still contribute reactive oxygen through the Fenton reaction. The accumulation of lipofuscin therefore has the potential to generate a self‐amplifying loop: more lipofuscin would mean more reactive oxygen, which would prevent a greater burden to proteolysis, which would mean the accumulation of more lipofuscin, and so on. At some point, the system would be expected to fail catastrophically, leaving the cell awash in its own toxic waste. An intriguing and poorly understood aspect of the garbage catastrophe is the effect proteasome inhibition has on lysosomal degradation and vice versa. The available evidence suggests that inhibition of either of the two major proteolytic systems will rapidly lead to inhibition of the other (Sitte et al., 2000; Terman and Sandberg, 2002; Sullivan et al., 2004) with the induction of a senescent phenotype in primary cell cultures (Terman and Brunk, 1998a; Chondrogianni and Gonos, 2004; Torres et al., 2006), despite the fact that the proteolytic systems exist within separate subcellular compartments. A facile explanation for these cross‐inhibitory effects would be that accumulated waste ‘‘spills over’’ into the other compartment. For example, it is conceivable that by occupying many proteasomes a large burden of cytosolic or nuclear waste would preclude the efficient functioning of the endoplasmic reticulum‐associated degradation pathway (ERAD) (Meusser et al., 2005), and that an ensuing increased burden of misfolded protein within the ER lumen might be relayed to the lysosome for degradation. Cross‐inhibitory scenarios could be postulated in reverse, originating within the membrane‐bound organelles and ultimately taxing proteasomal degradation, again intersecting at ERAD. These scenarios posit that it is the proteasome that is in limited supply, but a more nuanced hypothesis for the interdependency of the lysosomal and proteasomal degradation systems would place ubiquitin in the central role. In addition to its widely appreciated and well‐understood role in directing protein substrates to the proteasome, ubiquitin plays an important role in directing substrate traffic within the endosomal/lysosomal system (reviewed in d’Azzo et al. (2005)). If available ubiquitin pools are tied up in the proteasomal degradation of a bolus of misfolded or damaged proteins, the endosomal/lysosomal systems may operate at reduced capacity, leading to the accumulation of waste within the vesicular systems.

5 Genetic Models of Age‐Related Proteolytic Deficit The advent of gene targeting in rodents has provided the means by which causal relationships can at last be distinguished from mere correlation with regard to declining protease activity in the brain. In other words, while it is clear that certain activities decline with age, one must ask if the loss of specific protease activities sufficient to precipitate either neuronal dysfunctional or loss or both, or whether the reduction in proteolytic activity is symptomatic of a complex biological system in decline. The early indications from gene‐targeting experiments suggest that loss of protease function can in fact drive neurodegeneration, and that this approach has considerable promise in revealing the relative sensitivity of specific neuronal populations to proteolytic impairment. In rodents, genetically engineered deficiency in the lysosomal cathepsin proteases has had utility as models for neuronal ceroid lipofuscinoses (NCL), progressive and fatal neurodegenerative diseases affecting children. In these diseases, the accumulation of autofluorescent lipofuscin accompanies neuronal loss, resulting in seizures, loss of motor function, and ultimately death. The best characterized mouse model of NCL has a loss of function mutation not in a protease gene, but in the palmitoyl protein thioesterase‐1 (PPT‐1) gene that is a model of the CLN1 mutation in human NCL (Gupta et al., 2001) and whose deficiency leads to the accumulation of small fatty acylated peptides. Inactivation of the gene encoding cathepsin D (the major aspartate protease within lysosomes) induces lysosomal dysfunction culminating in neurodegeneration with hallmarks of NCL (Koike et al., 2000). Inactivation of either cathepsin B or cathepsin L (ubiquitous cysteine proteases) is insufficient to generate neuronal dysfunction (Deussing et al., 1998; Nakagawa et al., 1998; Roth et al., 2000) probably because of functional redundancy. Mice lacking both cathepsin B and cathepsin L die early in postnatal life with massive brain atrophy (Felbor et al., 2002).

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Recently, rodent experiments have been reported in which the targeted genes were not proteolytic components of the lysosome, but rather components involved in directing cell constituents to the lysosome through autophagy. Mice with engineered mutations in the Atg5 or Atg7 genes develop a progressive neurodegeneration (Hara et al., 2006; Komatsu et al., 2006), with formation of intracellular inclusion bodies typical of human diseases such as Parkinson’s disease or the expanded polyglutamine disorders (Huntington’s disease and the spinocerebellar ataxias). The formation of proteinaceous inclusions in these mice suggests that autophagy is an important mechanism of clearing the protein aggregates before they reach a critical size beyond which disassembly and degradation are no longer possible. Proteasome function was not found to be impaired in the Atg7 mice, despite the presence of ubiquitinated inclusions. These data argue that the inclusions per se are not inhibitory to proteasome function, consistent with a sequestration role of inclusions that may actually be neuroprotective (Gray, 2001; Arrasate et al., 2004; Bodner et al., 2006). With regard to the ‘‘garbage catastrophe’’ model described previously, the proteinaceous inclusions in Atg7‐deficient mice must at the very least be incapable of inhibiting both major proteolytic systems, perhaps because they are relatively inert and do not, for example, generate reactive oxygen. The proteasome is a complex structure built up from numerous protein subunits. The analysis of these subunits through gene targeting is still at an early stage, but insights are already accumulating as to the role of the proteasome in organismal homeostasis. Given the central role of ubiquitin‐mediated proteolysis in cellular functions as pivotal as cell‐cycle regulation, it would not be surprising if loss‐of‐function mutations in proteasome genes had highly deleterious effects early in development. This prediction is borne out by Mov34 mice in which retroviral insertional mutagenesis was found to have disrupted the non‐ATPase S12 (Rpn8) subunit of the 26S proteasome, which is required for development beyond midgestation (Gridley et al., 1990; Dubiel et al., 1995; Tsurumi et al., 1995). It is likely that conditional mutants will be required for loss‐of‐function studies of most proteasome subunits in vivo, but there are proteasome subunits with specialized functions whose absence is less likely to be lethal. The immunoproteasome is a specialized version of the proteasome that generates peptides of optimum length and composition for presentation as antigens on the surface of cells (reviewed in Kloetzel and Ossendorp (2004)). The alteration in protease activity is accomplished by substitution of protease subunits within the b ring. Mice in which one such subunit (LMP2) was inactivated by gene targeting have a deficiency in antigen processing, but such mice are viable (Van Kaer et al., 1994). Relative to wild‐type mice the LMP2‐knockout mice demonstrate significant deficiencies in multiple proteasome activities with increasing age, accompanied by an increase in the abundance of oxidized protein in the brain and liver (Ding et al., 2006), suggesting that loss of even a specialized subunit of the proteasome will have deleterious consequences for the cellular waste disposal systems. We await the generation of additional proteasome subunit mutations that will take advantage of recent methodologies allowing deletion of genetic information within specific tissues at specific stages of development. It would, for example, be informative to impair proteasome function selectively within postmitotic neurons of the mature CNS. An independent approach to genetically analyzing the role of an impaired UPS in aging is to study animals in which proteolysis is made inefficient through depletion of the ubiquitin pool. A technical obstacle to the direct targeting of ubiquitin genes is presented by their existence in multiple copies, but the effects of depletion of cellular ubiquitin pools can be studied in mice with spontaneous mutations of enzymes involved in ubiquitin recycling. The deubiquitinating enzyme Usp14 is a proteasome‐associated enzyme thought to be involved in the recovery of ubiquitin from substrates destined for proteolysis (Borodovsky et al., 2001; Hu et al., 2005). The axJ mouse strain has a spontaneous recessive mutation in the Usp14 gene resulting in the loss of Usp14 activity (Wilson et al., 2002). As a consequence of the inability of cells to salvage ubiquitin from ubiquitinated substrates the ubiquitin pools become depleted in axJ mice (Anderson et al., 2005). The pathological consequences of depleted ubiquitin pools are evident only in the central nervous system of axJ mice; animals homozygous for the mutation show loss of cerebellar Purkinje neurons accompanied by severe behavioral disorders including resting tremors and hind limb paralysis. Spontaneous mutation of a gene encoding a second, unrelated deubiquitinating enzyme (UCH‐L1) has been found to affect a distinct neuronal population, those of the gracile nucleus of the medulla oblongata (Saigoh et al., 1999). It appears that the UCH‐L1 enzyme is required for the maintenance of neuronal progenitor cell morphology (Sakurai et al., 2006) which may in turn relate to the maintenance of ubiquitin


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pools; by binding to monomeric ubiquitin, UCH‐L1 is reported to prevent ubiquitin degradation (Sakurai et al., 2006). The restriction of pathological findings to specific brain regions in mice with germ line mutations of deubiquitinating enzymes is consistent with the hypothesis that neuronal populations have variable vulnerability to ubiquitin pool deficiencies. This vulnerability may be dictated by both intrinsic factors and extrinsic factors. As an example of the former, ubiquitin is known to be involved in synaptic remodeling (Ehlers, 2003; Patrick et al., 2003; Yi and Ehlers, 2005; Patrick, 2006), and hence the availability of robust ubiquitin pools may be particularly important for neurons with extensive dendritic processes and high synaptic activity (this might explain the sensitivity of Purkinje neurons, for example, to perturbations in ubiquitin pools). It is visually evident from immunohistochemical analysis of ubiquitin in brain sections that subpopulations of neurons in the CNS differ in their ubiquitin pools (> Figure 23-2). In terms of extrinsic factors, it is reasonable to posit that neurons experiencing large or fluctuating stresses may have

. Figure 23-2 Immunohistochemical detection of ubiquitin in the hippocampal formation of an 18‐month‐old FVB/N strain mouse (counterstained with hematoxylin). Note the greater intensity of ubiquitin immunostaining in CA1 versus CA3 neurons and in neurons of the hilus (H) as opposed to those of the dentate gyrus (DG). The scale bar corresponds to 200 mm

greater demands for the degradation of damaged proteins. Neurons experiencing sudden stress with limited availability of ubiquitin pools may be more vulnerable than those with abundant ubiquitin or in a more benign environment.

6 Summary Although the two major proteolytic systems of the cell exist in separate subcellular compartments there may be interdependency arising from utilization of common components (ubiquitin, for example, required for vesicular sorting in one system and for proteasomal targeting of substrates in the other, or proteasomes, required for clearance of misfolded substrates from both membrane‐bound and cytosolic compartments). There is accumulating evidence that the proteolytic capacity of both systems declines with age. From the available in vivo evidence it is not clear whether the decline of one system precedes (and contributes to) the decline of the other, or whether both decline synchronously. In the context of the garbage catastrophe model, the relative timing of these events is of little import. The accumulation of proteinaceous waste that

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can be neither degraded nor diluted (through cell growth and division) and is itself a source of reactive oxygen stress becomes inexorable, and the decline of neuronal function inescapable. The prospect exists for pharmaceutical or gene‐based interventions to delay the age‐related decline in proteolysis, but the complexity of the proteolytic systems and their interrelationships (> Figure 23-3) clearly presents obstacles to the development of agents that can halt the process. This will be a war on several fronts.

. Figure 23-3 Interactions of the lysosomal and proteasomal degradation pathways. Some of the interactions indicated by arrows are currently hypothetical, but are mechanistically plausible. The inhibition of endosomal/lysosomal trafficking by depleted ubiquitin pools is one such example. Reciprocal inhibition of the two major proteolytic systems has been demonstrated (see text)

Acknowledgments The author is grateful to Dr. John Woulfe for the image appearing in > Figure 23-1, and to Mei Zhang for the immunohistochemical staining in > Figure 23-2. Mechanisms proposed in the chapter have been refined through serial discussions with Maria Tsirigotis, to whom the author is indebted. The author’s work is supported by the Institute of Aging, Canadian Institutes of Health Research.

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Nakagawa T, Roth W, Wong P, Nelson A, Farr A, et al. 1998. Cathepsin L: Critical role in Ii degradation and CD4 T cell selection in the thymus. Science 280: 450-453. Nakanishi H. 2003. Neuronal and microglial cathepsins in aging and age‐related diseases. Ageing Res Rev 2: 367-381. Nakanishi H, Amano T, Sastradipura DF, Yoshimine Y, Tsukuba T, et al. 1997. Increased expression of cathepsins E and D in neurons of the aged rat brain and their colocalization with lipofuscin and carboxy‐terminal fragments of Alzheimer amyloid precursor protein. J Neurochem 68: 739-749. Nakanishi H, Tominaga K, Amano T, Hirotsu I, Inoue T, et al. 1994. Age‐related changes in activities and localizations of cathepsins D, E, B, and L in the rat brain tissues. Exp Neurol 126: 119-128. Passmore LA, Barford D. 2004. Getting into position: The catalytic mechanisms of protein ubiquitylation. Biochem J 379: 513-525. Patrick GN. 2006. Synapse formation and plasticity: Recent insights from the perspective of the ubiquitin proteasome system. Curr Opin Neurobiol 16: 90-94. Patrick GN, Bingol B, Weld HA, Schuman EM. 2003. Ubiquitin‐mediated proteasome activity is required for agonist‐induced endocytosis of GluRs. Curr Biol 13: 2073-2081. Petropoulos I, Conconi M, Wang X, Hoenel B, Bregegere F, et al. 2000. Increase of oxidatively modified protein is associated with a decrease of proteasome activity and content in aging epidermal cells. J Gerontol A Biol Sci Med Sci 55: B220-B227. Ponnappan U, Zhong M, Trebilcock GU. 1999. Decreased proteasome‐mediated degradation in T cells from the elderly: A role in immune senescence. Cell Immunol 192: 167-174. Roth W, Deussing J, Botchkarev VA, Pauly‐Evers M, Saftig P, et al. 2000. Cathepsin L deficiency as molecular defect of furless: Hyperproliferation of keratinocytes and perturbation of hair follicle cycling. FASEB J 14: 2075-2086. Saigoh K, Wang YL, Suh JG, Yamanishi T, Sakai Y, et al. 1999. Intragenic deletion in the gene encoding ubiquitin carboxy‐ terminal hydrolase in gad mice. Nat Genet 23: 47-51. Sakurai M, Ayukawa K, Setsuie R, Nishikawa K, Hara Y, et al. 2006. Ubiquitin C‐terminal hydrolase L1 regulates the morphology of neural progenitor cells and modulates their differentiation. J Cell Sci 119: 162-171. Sheaff RJ, Singer JD, Swanger J, Smitherman M, Roberts JM, et al. 2000. Proteasomal turnover of p21Cip1 does not require p21Cip1 ubiquitination. Mol Cell 5: 403-410. Sitte N, Huber M, Grune T, Ladhoff A, Doecke WD, et al. 2000. Proteasome inhibition by lipofuscin/ceroid during postmitotic aging of fibroblasts. FASEB J 14: 1490-1498.

Sullivan PG, Dragicevic NB, Deng JH, Bai Y, Dimayuga E, et al. 2004. Proteasome inhibition alters neural mitochondrial homeostasis and mitochondria turnover. J Biol Chem 279: 20699-20707. Terman A. 1995. The effect of age on formation and elimination of autophagic vacuoles in mouse hepatocytes. Gerontology 41 Suppl 2: 319-326. Terman A. 2001. Garbage catastrophe theory of aging: Imperfect removal of oxidative damage? Redox Rep 6: 15-26. Terman A, Brunk UT. 1998a. Ceroid/lipofuscin formation in cultured human fibroblasts: The role of oxidative stress and lysosomal proteolysis. Mech Ageing Dev 104: 277-291. Terman A, Brunk UT. 1998b. Lipofuscin: Mechanisms of formation and increase with age. APMIS 106: 265-276. Terman A, Brunk UT. 2004. Lipofuscin. Int J Biochem Cell Biol 36: 1400-1404. Terman A, Sandberg S. 2002. Proteasome inhibition enhances lipofuscin formation. Ann N Y Acad Sci 973: 309-312. Tofaris GK, Layfield R, Spillantini MG. 2001. a‐Synuclein metabolism and aggregation is linked to ubiquitin‐ independent degradation by the proteasome. FEBS Lett 509: 22-26. Torres C, Lewis L, Cristofalo VJ. 2006. Proteasome inhibitors shorten replicative life span and induce a senescent‐like phenotype of human fibroblasts. J Cell Physiol 207: 845853. Tsurumi C, De Martino GN, Slaughter CA, Shimbara N, Tanaka K. 1995. cDNA cloning of p40, a regulatory subunit of the human 26S proteasome, and a homolog of the Mov‐ 34 gene product. Biochem Biophys Res Commun 210: 600608. Van Kaer L, Ashton‐Rickardt PG, Eichelberger M, Gaczynska M, Nagashima K, et al. 1994. Altered peptidase and viral‐ specific T cell response in LMP2 mutant mice. Immunity 1: 533-541. Viteri G, Carrard G, Birlouez‐Aragon I, Silva E, Friguet B. 2004. Age‐dependent protein modifications and declining proteasome activity in the human lens. Arch Biochem Biophys 427: 197-203. von Zglinicki T, Nilsson E, Docke WD, Brunk UT. 1995. Lipofuscin accumulation and ageing of fibroblasts. Gerontology 41 Suppl 2: 95-108. Wilson SM, Bhattacharyya B, Rachel RA, Coppola V, Tessarollo L, et al. 2002. Synaptic defects in ataxia mice result from a mutation in Usp14, encoding a ubiquitin‐ specific protease. Nat Genet 32: 420-425. Wolf DH, Hilt W. 2004. The proteasome: A proteolytic nanomachine of cell regulation and waste disposal. Biochim Biophys Acta 1695: 19-31. Yi JJ, Ehlers MD. 2005. Ubiquitin and protein turnover in synapse function. Neuron 47: 629-632.

24

From Concept to Potential Therapeutics: Neuroprotective Peptides

I. Gozes . J. Tiong

1 1.1 1.1.1 1.1.2 1.1.3 1.1.4

Neuroprotection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 674 Neuroprotective Proteins and Peptides: SNV, ADNF, and ADNP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 674 SNV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 675 The Discovery of ADNF and ADNP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 675 ADNF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 676 ADNP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 677

2 2.1 2.2 2.2.1 2.2.2 2.2.3 2.2.4 2.2.5 2.2.6 2.2.7 2.2.8 2.2.9

NAP and ADNF‐9 (SAL) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 678 In Vitro Neuroprotection by NAP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 678 In Vivo Neuroprotection by NAP and ADNF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 678 Cerebral Palsy Mouse Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 678 Apolipoprotein E Knockout Mouse Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 678 Fetal Alcohol Syndrome Mouse Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 678 AF64A Cholinotoxicity Rat Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 681 Traumatic Brain Injury Mouse Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 681 Permanent Midcerebral Artery Occlusion Rat Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 681 Memory in Middle‐Aged Rat Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 681 Anxiety Animal Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 682 Amyotrophic Lateral Sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 682

3 3.1 3.2

Mechanism of Neuroprotection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 682 Mechanism of Action of NAP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 682 Mechanism of Action of ADNF‐9/SAL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 683

4

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 684

#


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From concept to potential therapeutics: Neuroprotective peptides

Abstract: This chapter outlines the discovery and activity of the potent neuroprotective peptides ADNF‐9 (SAL) and NAP. These peptides are fragments of larger proteins, activity‐dependent neurotrophic factor (ADNF) and activity‐dependent neuroprotective protein (ADNP). The discovery of these proteins and peptides was a result of long‐term collaborative studies between Douglas E. Brenneman and Illana Gozes to reveal the mechanisms and molecules associated with neuropeptide‐based neuroprotection, particularly related to vasoactive intestinal peptide (VIP). Recent studies identified tubulin in glia (the brain support cells) as a target for NAP actions. It is suggested that NAP provides astrocyte protection by enhancing tubulin assembly into microtubules. These results paved the path to the initiation of clinical trials led by Allon Therapeutics, Inc., the Neuroprotection Company, with the lead compound NAP (formulations: AL‐108 and AL-208). List of Abbreviations: Aβ, β-amyloid peptide; ADNF, activity-dependent neurotrophic factor; ADNP, activity-dependent neuroprotective protein; ALS, amyotrophic lateral sclerosis; ApoE, apolipoprotein E; bFGF, basic fibroblast growth factor; BSO, Buthionine sulfoximine; CABG, cardiac artery bypass grafting; CaMKIV, Ca2+/calmodulin-dependent protein kinase IV; CNS, central nervous system; GSH, reduced glutathione; HIV, human immunodeficiency virus; hsp60, heat-shock protein 60; NMDA, N-methyl-Daspartate; PLAIDD, p75-like apoptosis-inducing death domain; P75NTR, p75 neurotrophin receptor; SNV, stearyl-Nle17-VIP; VIP, vasoactive intestinal peptide

1

Neuroprotection

There are many untreatable neurodegenerative diseases. Neurons are fundamental building blocks that underlie brain/nervous system functions including consciousness, memory, vision, speech, movement, as well as breathing, eating, and sleeping. Natural aging, neurodegenerative diseases, and traumatic injuries lead to the death of neurons, which are generally not replaced. The result includes cognitive decline, memory failure, loss of motor coordination, and ultimately death. Keeping the brain’s neurons alive under the many circumstances that lead to their demise is the critical goal of neuroprotection. Neuroprotection is the use of an agent that prevents neuronal death by blocking the pathophysiological steps resulting from insults and injuries. The field of neuroprotection has dramatically grown over the last decade due to advances in the understanding of the molecular mechanisms of cell death, disease pathology, and the absolute requirement for an effective neuroprotective therapy in humans. Neuroprotection is needed in chronic diseases such as Alzheimer’s disease, Parkinson’s disease, multiple sclerosis, and neuropathy, and acute disorders such as reperfusion injury postcardiac artery bypass grafting (CABG), stroke, and traumatic brain injury. Neuroprotection may be used to prevent progression of a chronic disease, administered following a central nervous system (CNS) trauma to inhibit secondary neuronal loss, or prior to surgical procedures known to cause cognitive impairment. Neuronal death is now understood to involve common molecular mechanisms across many of these diseases and insults: programmed cell death that is termed apoptosis. Therefore, it is likely that an agent that blocks fundamental steps in the mechanisms of apoptotic neuronal pathways will have multiple therapeutic applications. Studies have shown that endogenous neuroprotection occurs as a natural response of the CNS to injury and involves altered gene expression and production of neurotrophic factors. However, in cases when the brain is dying, endogenous mechanisms are not enough and external therapeutic intervention is required.

1.1 Neuroprotective Proteins and Peptides: SNV, ADNF, and ADNP Since the discovery of nerve growth factor, a consensus has emerged that neurotrophic proteins (Patterson, 1992; Maness et al., 1994) and neuropeptides (Gozes and Brenneman, 1993) have important regulatory functions during development and after nerve injury (Gozes et al., 1987; Brenneman et al., 1988; Gozes et al., 1996a, b, 1997a, b, c). Among the known neuroprotective peptides, vasoactive intestinal peptide (VIP) (Said and Mutt, 1970; Said, 1991) has been suggested to protect neurons against electrical blockade


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(Brenneman and Eiden, 1986; Brenneman et al., 1998a, b). Blockade of spontaneous electrical activity in spinal cord cell cultures results in the death of 50% of the neurons during a critical period of development. Furthermore, VIP is also identified as a modulator of growth, survival, and differentiation in many cell systems, including the brain, the gastrointestinal tract, lungs, and immune system, of both primary origin and cancerous ones (Gozes et al., 1999). Original and recent genetic manipulation to reduce VIP expression, or treatment with potent VIP antagonists resulted in diurnal rhythms defects as well as in developmental and behavioral deficits (Gozes et al., 1989, 1993, 1995; Glowa et al., 1992; Gressens et al., 1994; Wu et al., 1997; and for a recent review, see Gozes and Furman, 2004).

1.1.1 SNV Stearyl‐Nle17‐VIP (SNV) is a lipophilic superactive analog of VIP that contains two chemical modifications in VIP: the addition of an N‐terminal long‐chain aliphatic acid and the substitution of the Met in position 17 with Nle (Gozes and Fridkin, 1992; Gozes and Furman, 2004). These changes confer stability, longer half‐ life, and increased bioavailability. This analog exhibited both 100‐fold potency as compared with VIP (maximal effect manifested at 1 pM) and specificity for a VIP receptor in neuronal survival and neuroprotection against the b‐amyloid peptide fragment (the Alzheimer’s Disease neurotoxin) (Gozes et al., 1995, 1996a or b). It has been shown that VIP stimulates cGMP accumulation by several fold in isolated rat pinealocytes (Ho et al., 1987; Ogiwara et al., 1995; Schaad et al., 1995). Previous studies performed on astroglial cells revealed that VIP induced increases in cAMP formation (Magistretti and Schorderet, 1984) through a low‐ affinity receptor (Gozes et al., 1991), whereas SNV was inactive (Gozes et al., 1995). A cGMP antagonist (RP‐8‐pCPT‐cGMPS; 1012 to 109 M) reduced the number of surviving neurons (40–60%); this decline was spared in the presence of SNV (1013 M). A cGMP agonist (Sp‐8‐pCPT‐cGMPS; 1014 to 108 M) and SNV (1016 to 108 M) both provided significant neuroprotection against 1012 M of the cGMP antagonist. Immunoassays indicated that SNV induced increases in cGMP (2‐ to 3‐fold) in these cultures, whereas VIP was 1000‐fold less potent. These results implicate cGMP as a second messenger for VIP/SNV‐mediated effects on neuronal survival (Ashur‐Fabian et al., 1999). The developmental and protective effects of SNV were also investigated in vivo using two models of developmental retardation, hypoxia and cholinergic blockade. In both cases, chronic administration of SNV during development provided protective effects. Water maze experiments on weaned animals have demonstrated a prophylactic action for SNV and enhancement of spatial memory in animals exposed to a cholinotoxin. SNV may act by providing neuroprotection, thereby improving cognitive functions (Gozes et al., 1996a, b, 1997a, b, c, 1998). In a model of cerebral palsy, SNV showed brain protection, even after peripheral administration (Gressens et al., 1999).

1.1.2 The Discovery of ADNF and ADNP VIP prevents cell death that is associated with electrical blockade by releasing glial‐derived, survival‐ promoting substances (Brenneman and Gozes, 1996; Gozes and Brenneman, 1996). Previous studies have identified several components within the neurotrophic milieu produced by VIP‐stimulated astroglia, the most potent being activity‐dependent neurotrophic factor (ADNF) (Brenneman and Gozes, 1996; Gozes and Brenneman, 1996; Gozes et al., 1997c; Brenneman et al., 1998a). Comparative studies with other recognized growth factors indicated that at femtomolar concentration only ADNF prevented neuronal cell death associated with electrical blockade in rat cerebral cortical cultures (Gozes et al., 1997c). A 14‐amino‐ acid peptide derived from ADNF (ADNF‐14, VLGGGSALLRSIPA) has been reported to protect cultured neurons from multiple neurotoxins (Brenneman and Gozes, 1996). This ADNF‐derived peptide exhibited a remarkable structural similarity to the stress‐induced chaperonin, heat‐shock protein 60 (hsp60), containing the highly conserved sequence VLGGGCALLRCIPA (Gozes and Brenneman, 1996). Structure–activity relationships of peptides related to ADNF‐14 revealed a nine‐amino‐acid core peptide (ADNF‐9,

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also termed SAL: SALLRSIPA) with greater potency and broader effective concentration range (10–16 to 10–13 M) than ADNF or ADNF‐14 in preventing neuronal death associated with tetrodotoxin treatment (Brenneman et al., 1998b). When compared with other neuroprotective protein factors such as humanin, insulin‐like growth factor I, and basic fibroblast growth factor (bFGF), the ADNF peptide protected against toxicity associated with b‐amyloid peptide at a higher potency (Hashimoto et al., 2004). Antibodies prepared against ADNF‐14 and affinity‐purified to recognize ADNF‐9 (SAL) were used to screen a cDNA expression library of neuroglial origin. A novel ADNF‐14/9‐like active peptide (NAPVSIPQ or NAP) with a greater in vivo neuroprotective efficacy, compared with ADNF‐9 (SAL), was discovered that constitutes a part of a new protein. As the cloned protein contained the potent peptide that prevented neuronal death associated with electrical blockade, it was named the activity‐dependent neuroprotective protein (ADNP) (Bassan et al., 1999).

1.1.3 ADNF Amino acid sequencing of ADNF peptide fragments indicated homology to hsp60 (Brenneman and Gozes, 1996; Gozes and Brenneman, 1996). Antibodies to ADNF recognized a peptide sequence in ADNF (VLGGGSALLRSIPA, ADNF‐14) that is homologous to hsp60 (VLGGGCALLRCIPA, hsp60‐14) (Gozes and Brenneman, 1996), however, only minimal cross‐reactivity was observed with hsp60. Treatment of cerebral cortical neurons with this antiserum increased the number of apoptotic neurons and cotreatment with ADNF prevented the antibody‐induced death. Unexpectedly, ADNF‐14 was found to mimic the survival‐promoting activity of ADNF (Brenneman and Gozes, 1996). By comparison, hsp60‐14, was 105‐fold less potent than ADNF‐14 and less efficacious (Brenneman and Gozes, 1996). ADNF‐14 mimics a reduced form of hsp60, whereas hsp60‐14 may be oxidized, exhibiting a different structure. Along the same line of thought, antibodies against hsp60, recognizing the reduced protein, produced neuronal cell death that was inhibited by ADNF, but not by native hsp60 (Brenneman and Gozes, 1996). Hsp60, localized in widely divergent cellular compartments, including mitochondria, endoplasmic reticulum, peroxisomes, granules, and plasma membrane, has been assigned a chaperonin function not only in the mitochondria, but also in the plasma membrane, regulated by protein kinase A (Khan et al., 1998). Furthermore, hsp60 secretion into the extracellular milieu of neurons and glia increased following a mild stress and/or exposure to VIP (Bassan et al., 1998). The question of whether hsp60 is processed to yield ADNF or whether ADNF is a different gene product presents a subject for future research. Recent experiments indicate that prevention of hsp60 expression by antisense oligodeoxynucleotides results in decreased ADNF‐like immunoreactivity and increasing hsp60 expression by cDNA transfection results in increased ADNF‐like expression. These data suggest an association between ADNF and hsp60 expression (Gozes et al., 2005a). ADNF‐Derived Peptides: Structure–Activity Relations Regardless of the hsp60/ADNF relation, structure‐ activity experiments were further conducted in search of the essential ADNF amino‐acid sequence required for neuroprotection. Screening of 40 peptides identified SALLRSIPA (ADNF‐9 ¼ SAL), a peptide that captured and exceeded the neurotrophic properties of ADNF‐14 and the parent molecule ADNF (Brenneman and Gozes, 1996; Brenneman et al., 1998b). The increased range of effective concentrations obtained by peptide shortening may be attributed to the requirement of the specific SALLRSIPA epitope for neuroprotection, and structural constraints of the larger molecules. Further studies identified an ADNF‐like peptide termed NAP (NAPVSIPQ) (Bassan et al., 1999) and structure activity studies identified the shared SIP epitope as important for biological activity (Wilkemeyer et al., 2003). Additional structural/functional experiments showed that the ADNF‐9 activity is not chiral and substitution of all the ADNF‐9 (SAL) L‐amino acids by D‐amino acids resulted in biologically active peptides that offer increased stability (Brenneman et al., 2004; Wilkemeyer et al., 2004). Furthermore, Chiba et al., found protective activity against neurotoxicity in a mouse model of familial amyotrophic


24

lateral sclerosis (ALS) also in ALLRSIPA, although it was a 100‐fold less potent than SALLRSIPA ¼ ADNF‐ 9/SAL (Chiba et al., 2004). For more information on ADNF please consult a recent review (Gozes et al., 2005a).

1.1.4 ADNP ADNP (Bassan et al., 1999) was also identified as a VIP‐ and as VIP‐related peptides responsive gene during brain development (Bassan et al., 1999; Pinhasov et al., 2003; Zusev and Gozes, 2004). VIP has dramatic effects on embryonic growth at the time of neural tube closure (E8.5–9) (Pinhasov et al., 2003) and affects brain development and function (Gozes and Brenneman, 1989; Gozes et al., 1993; Gressens et al., 1993). VIP provides a neuroprotective milieu (Brenneman and Eiden, 1986) by activating glial cells. VIP expression is developmentally determined (Gozes et al., 1988), increases with brain maturation (Gozes et al., 1987), stimulates synapse formation (Blondel et al., 2000) and decreases with aging (Gozes et al., 1988). It is our hypothesis that some of the activities attributed to VIP are implemented via the activation of ADNP and downstream pathways. The ADNP gene is highly conserved in the mouse (Bassan et al., 1999), human (Zamostiano et al., 2001), and rat (Sigalov et al., 2000), and abundantly expressed in the brain and in other tissues (Bassan et al., 1999). Pronounced ADNP mRNA expression in the hippocampus, cerebral cortex, and cerebellum (Bassan et al., 1999) suggests an involvement for the protein in brain metabolism. Recent studies with knockout embryos identified ADNP as a protein essential for brain formation and embryonic development (Pinhasov et al., 2003). The deduced protein structure of ADNP contains a homeobox domain profile that includes a nuclear export signal indicating that ADNP may have nuclear and extracellular functions (Gozes et al., 2000b). Another sequence motif found in ADNP is a cytoplasmic import sequence, suggesting internalization into target cells (Furman et al., 2004). ADNP also contains a small eight‐amino‐acid peptide sequence termed NAP (NAPVSIPQ), a motif identified by peptide scanning, that protects neurons, at femtomolar concentrations, against a wide variety of toxic substances in vitro and in vivo in models of neuronal injury (Bassan et al., 1999; Gozes and Brenneman, 2000; Gozes et al., 2000a; Beni‐Adani et al., 2001; Leker et al., 2002). Utilizing antibodies directed against the NAP epitope, ADNP‐like immunoreactivity was found in both the cytoplasmic and in the nuclear cell fractions of astrocytes. In the cytoplasm, ADNP‐like immunoreactivity was colocalized with tubulin‐like immunoreactivity and with microtubular structures, but not with actin microfilaments. Since microtubules are key components of the developing neuron and the developing brain, the possible tubulin–ADNP interaction may suggest a functional correlate to ADNP’s role in the brain. In addition, ADNP‐like immunoreactivity was detected in the extracellular milieu of astrocytes, and its content was increased by 1.4‐fold after incubation of the astrocytes with VIP. VIP is known to activate astrocytes to secrete neuroprotective/neurotrophic factors, and it is now suggested that ADNP constitutes part of this VIP‐ stimulated protective milieu (Furman et al., 2004). Gene expression studies have shown that ADNP is an injury‐protection‐responsive gene, increasing in pheochromocytoma (neuronal‐like cellular model) upon treatment with SNV in response to protection against ischemic stress (Sigalov et al., 2000). Furthermore, in head trauma models, ADNP expression changes in a time‐dependent manner with potential decreases close to the time of injury and long‐term increases in astrocytic‐like cells that may represent part of a compensatory endogenous neuroprotection mechanism (Gozes et al., 2005b; Zaltzman et al., 2004). These changes in ADNP expression are regulated, in part, by the genetic background of the individual subject, with the proinflammatory Mac‐1 knockout mice showing increased resistance to the adverse consequences of head trauma and no increases in long‐term ADNP gene expression (Zaltzman et al., 2005). It is hypothesized that ADNP is a development/plasticity/injury‐associated gene. ADNP is highly expressed in the embryonic nervous system (Poggi et al., 2002; Pinhasov et al., 2003) and is responsive to stress conditions in the developing embryo (Poggi et al., 2003). In the mature brain (adult mouse hypothalamus), ADNP expression is sex dependent, with increased expression in the arcuate nucleus that exhibits estrus cycle‐associated rhythmicity (Furman et al., 2005).

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NAP and ADNF‐9 (SAL)

2.1 In Vitro Neuroprotection by NAP Neuroprotective activities of NAP have been investigated in numerous in vitro systems following the application of neurotoxic compounds and procedures such as: the Alzheimer’s disease toxin (b‐amyloid peptide or Ab), the toxic envelope protein of the human immunodeficiency virus (HIV; gp120), glucose deprivation, electrical blockade (tetrodotoxin), oxidative stress (hydrogen peroxide and glutathione), dopamine toxicity, and excitotoxicity (N-methyl-D-aspartate) (Bassan et al., 1999; Offen et al., 2000; Steingart et al., 2000; Zemlyak et al., 2000; Lagreze et al., 2005). These studies indicated an exceptionally broad range of neuroprotective efficacy against these toxic insults. In addition, human cortical neurons isolated from brain tissue of patients with Down’s syndrome are also protected by NAP when exposed to oxidative stress induced by excess levels of Ab (Busciglio et al., In press). Additional studies suggest that NAP stimulates axonal outgrowth, a process that precedes synapse formation (Lagreze et al., 2005; Smith‐Swintosky et al., 2005). Together, these in vitro data provide evidence that NAP may be effective as a treatment against Alzheimer’s disease‐associated cell death. A summary of the in vitro pharmacology of NAP is listed in > Table 24-1.

2.2 In Vivo Neuroprotection by NAP and ADNF 2.2.1 Cerebral Palsy Mouse Model A potential model for brain lesions associated with cerebral palsy is the cerebral palsy mouse model. Here, ibotenate excitotoxicity in newborn and 5‐day‐old mice was evaluated. Intracerebral injections of ADNF‐14 (10 pg) provided significant protection against ibotenate‐induced microgyric‐like cortical lesions and white matter cysts (Gressens et al., 1997).

2.2.2 Apolipoprotein E Knockout Mouse Model The E4 allele of the gene encoding the lipid carrier apolipoprotein E (ApoE) is a risk factor for Alzheimer’s disease. ApoE‐deficient mice are developmentally retarded and exhibit short‐term memory impairments (Gozes et al., 1997a). Daily subcutaneous injections of ADNF‐9/SAL or NAP (mg amounts, for the first 2 weeks of life) to newborn apoE‐deficient mice, accelerated the acquisition of developmental reflexes (cliff drop aversion and limb placing) (Bassan et al., 1999). The acceleration was more pronounced with NAP as compared with ADNF‐9/SAL > ADNF‐14. NAP treatment (and not ADNF‐14 or ADNF‐9/SAL treatment) also provided significant increases in brain choline acetyl transferase activity and prevented short‐term memory deficits (measured 1 week after cessation of peptide injection) (Bassan et al., 1999). The increased efficacy of NAP may be associated with enhanced stability and bioavailability as compared with the larger ADNF peptides (Brenneman et al., 1998b). In this study, drug delivery to the brain was facilitated by the fact that the blood–brain barrier is not yet established during the initial postnatal development of the rodent. A summary of the in vivo pharmacology of NAP is listed in > Table 24-2.

2.2.3 Fetal Alcohol Syndrome Mouse Model NAP and ADNF‐9/SAL have been shown to provide protective intervention in a model of fetal alcohol syndrome. Fetal demise and growth restrictions were produced after intraperitoneal injection of ethanol to pregnant mice during midgestation (E8) (Spong et al., 2001). Death and growth abnormalities elicited by alcohol treatment during development are believed to be associated, in part, with severe oxidative damage. NAP and ADNF‐9/SAL have been shown to exhibit antioxidative and antiapoptotic actions in vitro.


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. Table 24-1 Summary of in vitro pharmacology of NAP Insult Oxidative stress (H2O2)

Reduced glutathione (GSH) Buthionine sulfoximine (BSO) Electrical blockade (tetrodotoxin) Alzheimer’s disease related toxicity (b amyloid peptide fragments)

Excitotoxicity (N‐methyl‐D‐ aspartate, NMDA) AIDS‐related toxicity (gp120, envelope protein of the virus) Down’s syndrome Glucose depravation Dopamine toxicity Ethanol toxicity

Serum‐free conditions Tumor necrosis a toxicity Zinc toxicity

Model Rat pheochromocytoma (PC12 cells) Human embryonic cortical neurons Human neuroblastoma

Rat cerebral cortical neurons in mixed neuronal–glial cultures Rat cerebral cortical neurons in mixed neuronal‐ glial cultures Rat cerebral cortical neuron enriched cultures Rat cerebral cortical neurons in mixed neuronal–glial cultures Rat cerebral cortical neurons in mixed neuronal–glial cultures Human embryonic cortical neurons (abortion) Rat cerebral cortical neuron‐enriched cultures Rat pheochromocytoma (PC12 cells) Alcohol sensitive, human L1‐transfected NIH/3T3 cells Newborn (P0–P2) rat retinal ganglion cells Rat pheochromocytoma (PC12 cells) Rat cortical astrocytes

Protective concentrations 10–17 to 10–14 M

Reference Steingart et al. (2000)

10–15 to 10–13 M

Busciglio et al. (In press)

10–14 to 10–13 M; 10–10 to 10–7 M

Offen et al. (2000)

10–18 to 10–14 M; 10–11 to 10–9 M 10–16 to 10–15 M

Bassan et al. (1999); Brenneman et al. (2000); Wilkemeyer et al. (2003) Bassan et al. (1999)

10–13 to 10–11 M

Zemlyak et al. (2000)

10–16 to 10–8 M

Bassan et al. (1999)

10–15 to 10–10 M

Bassan et al. (1999)

10–15 M

Busciglio et al. (In press)

10–12 M

Zemlyak et al. (2000)

10–18 to 10–10 M

Offen et al. (2000)

≧10–16 M

Wilkemeyer et al. (2002)

10–14 to 10–10 M

Lagreze et al. (2005)

10–14 M

Beni‐Adani et al. (2001)

10–15 to 10–10 M

Divinski et al. (2004)

Pretreatment with an equimolar combination of the peptides prevented the alcohol‐induced fetal death and growth abnormalities. Pretreatment with NAP alone resulted in a significant decrease in alcohol‐associated fetal death, whereas ADNF‐9/SAL alone had no detectable effect on fetal survival after alcohol exposure, indicating a pharmacological distinction between the peptides. Biochemical assessment of the fetuses indicated that the combination peptide treatment prevented the alcohol‐induced decreases in reduced glutathione. Peptide efficacy was evident with either 30‐min pretreatment or with 1‐h postalcohol administration. Bioavailability studies with [3H]NAPVSIPQ indicated that 39% of the total radioactivity comigrated with intact peptide in the fetus 60 min after administration. These studies demonstrate that fetal death and growth restriction

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. Table 24-2 Summary of in vivo pharmacology of NAP Insult Cholinotoxicity

Model Rats

Apolipoprotein E deficiency (gene knockout) Head trauma

Mice

Mice

Mice

Administration Chronic intranasal Chronic (developmental) subcutaneous Acute subcutaneous

Chronic (developmental) subcutaneous Acute intravenous

Protective concentrations 2 mg/kg

Outcome Cognitive enhancement Cognitive enhancement

Reference Gozes et al. (2000a) Bassan et al. (1999)

0.25–0.3 mg/g

Reduced mortality and enhanced clinical recovery

0.25–0.5 mg/g

Enhanced clinical recovery and cognition Protection against apoptosis and enhanced clinical recovery Protection against paralysis

Beni‐Adani et al. (2001); Romano et al. (2002) Zaltzman et al. (2004)

0.25–0.5 mg/g

3, 30 mg/kg

Stroke (middle cerebral artery occlusion)

Hypertensive rats

Multiple sclerosis (autoimmune encephalomyelitis)

Mice

Chronic intranasal

20 mg/kg

Mice

Acute intravenous

0.3 mg/g

Mice

Acute intraperitoneal Chronic intranasal Chronic intranasal

1–2 mg/g

Fetal alcohol syndrome Middle age

Rats

Middle age

Mice

2 mg/kg 12.5mg/kg

Protection against paralysis and axonal damage Protection against fetal demise Cognitive enhancement Anxiolytic

Leker et al. (2002)

Gozes et al. (2003)

Spong et al. (2001) Gozes et al. (2002) Alcalay et al. (2004)

associated with prenatal alcohol exposure were prevented by combinatorial peptide treatment and suggest that this therapeutic strategy might be explored in other models/diseases associated with oxidative stress (Spong et al., 2001). A further study investigated ADNF‐9/SAL for its protective properties against fetal alcohol‐related brain growth retardation, using an established liquid diet model of alcohol‐related neurodevelopmental disorder in C57BL/6 mice (Zhou et al., 2004). Alcohol exposure during neurulation reduced body weight, head size, brain weight, and volume. Major gross brain deficits include restricted midline neural tissue growth leading to openings at the roof /floor plate. ADNF‐9/SAL treatment increased the fetal body weight, restored brain weight, brain volume, and regional brain size. Furthermore, ADNF‐9/SAL‐treatment restored cortical thickness, reduced the size and frequency of neural tube openings, and attenuated ventricular enlargement. The ability of ADNF‐9/SAL to antagonize alcohol‐retarded brain growth and development of forebrain and midline neural tube at midgestation suggests its potential use as an antagonist against fetal alcohol‐rendered microencephaly early in development (Zhou et al., 2004). These experiments are in agreement with studies performed by Michael Charness et al. (Wilkemeyer et al., 2002, 2003, 2004) showing that ADNF‐9/SAL and NAP may antagonize the adverse effects of alcohol on cell adhesion, effects that are of high importance during embryonic development. Interestingly, ADNP has been shown to be essential for neural tube closure


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(Pinhasov et al., 2003) and here the ADNF/ADNP peptides (i.e., ADNF‐9/SAL) are suggested to enhance neural tube closure.

2.2.4 AF64A Cholinotoxicity Rat Model In the rat cholinotoxicity model, acetylcholine producing neurons which are some of the earliest to undergo degeneration in Alzheimer’s disease are selectively killed following injection of the toxin AF64A (ethylcholine aziridium). Intranasal administration of NAP (2mg/kg/day) provided significant improvement in short‐term spatial memory measured in the Morris water maze (Gozes et al., 2000a). Long term cognitive enhancement and neuroprotective effects were also observed in the NAP‐treated animals compared with placebo controls (Gozes et al., 2000a, 2002).

2.2.5 Traumatic Brain Injury Mouse Model NAP was shown to be neuroprotective in a mouse model of closed head injury. NAP injection after injury reduced mortality and facilitated neurobehavioral recovery (P 0.005). Edema was reduced by 70% in the NAP‐treated mice (P 0.01). Furthermore, in vivo magnetic resonance imaging demonstrated significant brain tissue recovery in the NAP‐treated animals. NAP treatment decreased tumor necrosis factor‐a levels in the injured brain and was shown to protect pheochromocytoma (PC12 cells) against tumor necrosis factor‐a‐induced toxicity. Thus, NAP provides significant amelioration from the complex array of injuries elicited by head trauma (Beni‐Adani et al., 2001). Follow‐up studies have shown that NAP treatment may be prophylactic, thus, treatment during the first weeks of life resulted in long‐term neuroprotection that was manifested at several months of age, in faster recuperation from traumatic head injury (Zaltzman et al., 2004). These findings were associated with ADNP (Gozes et al., 2005b; Zaltzman et al., 2004) and NAP‐related changes in gene expression patterns after traumatic head injury. These trauma‐related changes include attenuated increases in the long‐term expression of the proinflammatory Mac‐1 (Romano et al., 2002; Zaltzman et al., 2004) and modified NAP‐ related protection time line in Mac‐1‐deficient mice that show partial protection against the adverse effects of traumatic head injury (Zaltzman et al., 2005).

2.2.6 Permanent Midcerebral Artery Occlusion Rat Model Leker et al. (2002) demonstrated that a single postinjury (up to 4 h) injection of NAP had durable protective effects (infarct size, sensory and motor reflexes) in a model of focal irreversible cerebral ischemia (Leker et al., 2002). NAP appeared to produce cerebroprotection by reducing apoptosis. NAP significantly reduced motor disability and infarct volumes compared with vehicle or D‐NAP when tested at 24 h after stroke onset (infarct volume 9.67 1.4% versus 17.04 1.18% and 19.19 1.9% of hemispheric volume, respectively; P 0.05). NAP given 4h but not 6 h after permanent middle cerebral artery occlusion still conferred significant neuroprotection (infarct volume 10.9 3.9% of hemispheric volume; P 0.05 versus vehicle). Long‐term studies demonstrated that infarct volumes and disability scores remained significantly lower after 30 days in NAP‐treated animals. NAP significantly reduced the number of apoptotic cells. The results indicate that the durable cerebroprotection mediated by NAP involves antiapoptotic mechanisms.

2.2.7 Memory in Middle‐Aged Rat Models The memory‐enhancing activity of NAP was evaluated in normal middle‐aged animals to further assess NAP’s breadth of neuroprotection. NAP was administered by nasal application. Results showed that in the paradigm of the Morris water maze, assessing short‐term memory, only the NAP‐treated middle‐aged rats

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and not placebo‐treated rats showed significant improvements by the end of the testing period. These results suggest efficacy for NAP in normal aging that is associated with accumulating environmental and genetic toxic factors (Gozes et al., 2002).

2.2.8 Anxiety Animal Model As indicated above, acute NAP administration by the intranasal route resulted in improved performance in the Morris water maze of normal and cognitively impaired rats. In these animals, it was observed, but not quantified, that NAP exhibited an anxiolytic effect (Gozes et al., 2002). Therefore, the effects of chronic NAP treatment on anxiety‐like behavior in mice in the elevated plus maze were evaluated. Results showed that 5 months of daily (intranasal) treatment with NAP reduced anxiety, measured as the percentage of time spent in the open arms of the maze (P 0.01). This effect was maintained after a longer (8 months) exposure to NAP. In addition, after 8 months of NAP treatment, the percentage of open arm entries out of total arm entries was significantly higher in the treated mice (P 0.01). Motor function indices indicated no significant differences between the groups. Furthermore, prolonged treatment with NAP (7 months) showed some beneficial effects on Morris water maze performance in the aging mice. It is concluded that NAP offers a unique combination of anxiolytic‐ and cognitive‐enhancing properties observed after prolonged chronic intranasal treatment (Alcalay et al., 2004).

2.2.9 Amyotrophic Lateral Sclerosis In vivo studies showed that intracerebroventricularly administered ADNF‐9/SAL significantly improved motor performance of G93A‐SOD1 (Cu/Zn‐superoxide dismutase‐1) transgenic mice, a widely used model of familial ALS, although survival was extended only marginally. Thus, the neuroprotective activity of ADNF‐9 provides a novel insight into the development of curative drugs for ALS (Chiba et al., 2004).

3

Mechanism of Neuroprotection

3.1 Mechanism of Action of NAP NAP is an eight‐amino‐acid peptide with the following structure: Asn‐Ala‐Pro‐Val‐Ser‐Ile‐Pro‐Gln (single letter code: NAPVSIPQ). This sequence exhibits the following structure–function characteristics: It has a lipophilic structure that allows penetration through lipid membranes as in the case of the cellular membrane and the blood–brain barrier (Divinski et al., 2004). It has intrinsic b sheet breaker characteristics, thus acting as a peptide chaperone to protect against toxic Ab plaque associated with Alzheimer’s disease (Ashur‐Fabian et al., 2003). It chelates trace amounts of heavy metals, thus preventing toxicity (Ashur‐Fabian et al., 2003). It binds to tubulin and promotes proper microtubule assembly—as a peptide chaperone (Divinski et al., 2004; Gozes and Divinski, 2004). NAP has demonstrated efficacy against the two primary Alzheimer’s disease pathologies—amyloid plaques and hyperphosphorylated tau (Bassan et al., 1999; Zemlyak et al., 2000; Ashur‐Fabian et al., 2003; Gozes and Divinski, 2004). In vitro experiments have demonstrated that the presence of NAP prevents the aggregation of Ab into amyloid plaques. In addition, the presence of NAP also promotes the dissolution of these plaques in cell culture and protects neuronal cells against the toxicity of Ab (Bassan et al., 1999; Zemlyak et al., 2000; Ashur‐Fabian et al., 2003). These findings are associated with the b sheet breaker characteristics of NAP as well as its ability to bind heavy metals that have been shown to promote Ab aggregation (Ashur‐Fabian et al., 2003).


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It is well recognized that tau performs a critical function of stabilizing and maintaining the microtubular network, which in turn is essential for axonal transport in neurons. The formation of neurofibrillary tangles, a hallmark of Alzheimer’s disease, which results from the hyperphosphorylation of tau, leads to microtubule breakdown and impaired axonal transport (Ishihara et al., 1999; Lee et al., 2001; Morfini et al., 2002). Furthermore, electron microscopic studies of brain tissue from patients with Alzheimer’s disease have shown a decrease in microtubules as compared with normal brains (Cash et al., 2003). Divinski et al. (2004) have demonstrated that exposure to zinc toxicity resulted in microtubule breakdown in astrocytes and that NAP protects these cells from this toxicity by promoting the reorganization of the microtubular network. In the same experiments, tubulin was identified as a NAP‐binding molecule (Divinski et al., 2004). Furthermore, in the presence of NAP, there is an increase in the ratio of nonphosphorylated tau to phosphorylated tau (Gozes and Divinski, 2004) and increased neurite outgrowth, a process that is dependent on slow axoplasmic transport (Gozes and Littauer, 1982; Lagreze et al., 2005; Smith‐Swintosky et al., 2005). Therefore, it is possible that NAP functions to promote the assembly and stability of the microtubular network either directly by binding to tubulin or indirectly through changes in the levels of the different forms of tau. These neuroprotective mechanisms increase the internal defense mechanisms of the cell (Ashur‐Fabian et al., 1999). Previous studies have suggested that deposition of Ab enhances tau phosphorylation in neuronal cultures (Busciglio et al., 1995; Ferreira et al., 1997) and in vivo when Ab is deposited in the brain (Sigurdsson et al., 1997; Geula et al., 1998; Ashur‐Fabian et al., 2003). Ab42 aggregates into oligomers along microtubules of neuronal processes, both in Tg2576 and human Alzheimer’s disease brain (Takahashi et al., 2004). Furthermore, there is evidence indicating that in the presence of Ab there is significant microtubule disruption. Michaelis et al. demonstrated that microtubule‐stabilizing agents have protective effects on microtubule structure in the presence of Ab (Michaelis et al., 2004). These results suggest that there may be a cause–effect relationship between the two distinct pathologies of Alzheimer’s disease and that NAP provides a neuroprotective effect through microtubule reorganization, thus allowing for the maintenance and stability of the cytoskeletal structure, which is altered by neurodegenerative insults.

3.2 Mechanism of Action of ADNF‐9/SAL Kinetic studies indicated that >2 h treatment with ADNF‐9/SAL produced neuroprotection for 4 days (Brenneman et al., 1998b). The immediate response of neurons to ADNF‐9/SAL may be attributed to effects on gene expression. Exposure to the Ab peptide resulted in decreased expression of hsp60, while 3 h cotreatment with ADNF‐9/SAL induced increases in neuronal hsp60 mRNA, providing an intracellular, neuroprotective milieu of chaperonins (Zamostiano et al., 1999). Prevention of receptor‐mediated endocytosis (30‐min bafilomycin A1 exposure) blocked the neuroprotective properties of ADNF‐9/SAL (Brenneman et al., 1998b). These studies suggest: (1) receptor‐mediated mechanism; (2) active uptake; and (3) intracellular receptors. The dose–response neuroprotection curve to ADNF‐9/SAL against TTX toxicity had a 10,000‐fold higher potency when glial cells are present, with attenuation at greater than picomolar concentrations. In contrast, in the absence of glia (with neurons plated on poly‐L‐lysine instead of on an astrocytic bed), attenuation was not observed (Brenneman et al., 1998b). This implies a glial role beyond providing ADNF or an ADNF precursor. The in vitro efficacy of the ADNF family against a multiplicity of neurotoxins was determined (Brenneman and Gozes, 1996; Brenneman et al., 1998a, b; Bassan et al., 1999; Guo et al., 1999). Neuroprotection was obtained against the disease‐relevant toxins, gp120 (neurotoxic envelope protein of the human immunodeficiency virus (HIV)), N‐methyl‐D‐aspartate (NMDA, excitotoxicity), and the Ab peptide. In hippocampal neurons from mutant presinilin‐1 knockin mice (associated with overexpression of the toxic Ab peptide and early onset Alzheimer’s disease) (Guo et al., 1999), a 24‐h pretreatment with 0.1 pM ADNF‐9/SAL or 100 ng/mL bFGF prior to exposure to glutamate excitotoxicity resulted in reduced oxyradical production and increased mitochondrial function, providing significant protection to both wild‐type and the more susceptible knockin neurons (Guo et al., 1999). Furthermore, the Ca2þ influx response to glutamate was suppressed in neurons pretreated with

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ADNF‐9/SAL and bFGF (Guo et al., 1999). This study places ADNF‐9/SAL on par with bFGF, both factors interrupting excitotoxic neurodegenerative cascades promoted by the presenilin‐1 mutation. As indicated above, recent studies performed by Hashimoto et al.(2004) suggest an increased potency for ADNF‐9/SAL compared with bFGF and other neurotrophic factors. Previous studies have shown that the Ab peptide binds to the p75 neurotrophin receptor (P75NTR) and to its relative p75‐like apoptosis‐inducing death domain (PLAIDD) (Hashimoto et al., 2004). The death mechanism induced by the Ab peptide includes activation of G0/Gi, Jnk, NADPH oxidase, and caspase‐3‐related caspases. ADNF‐9/SAL and other neurotrophins inhibit these pathways, with ADNF‐like molecules exhibiting a very high potency (Hashimoto et al., 2004). Interestingly, NAP was shown to inhibit caspase‐3 activation in vivo in a stroke‐like model in rats (Leker et al., 2002). Furthermore and as reviewed before, ADNF‐9 neuroprotection is associated with changes in gene expression (Gozes et al., 2005a) that may be related to CREB activation (White et al., 2000) and protein phosphorylation (protein kinase C and mitogen‐associated protein kinase kinase) (Gressens et al., 1999). Other results indicated association with certain tyrosine kinases and Ca2þ/calmodulin‐ dependent protein kinase IV (CaMKIV) (Chiba et al., 2004).

4

Conclusion

The mechanism of femtomolar neuroprotection is of high interest as the use of low doses should reduce or abolish side effects. Toxicological studies indicated that NAP presents safe drug profile and is suitable for clinical development. Allon Therapeutics licensed these neuroprotective peptides from the National Institute of Health and Ramot at Tel Aviv University. Given the increased efficiency of NAP in vivo neuroprotection, NAP (intranasal formulation, AL‐108) was chosen as a lead candidate for Alzheimer’s disease drug development. Allon has completed phase Ia human clinical trial evaluating the safety and pharmacokinetics of NAP in healthy adults. NAP was administered intranasally to healthy adults in a double‐blind, placebo‐controlled, randomized, sequential and ascending study to evaluate primarily the safety and pharmacokinetic results. The dosing was well tolerated by all subjects and no significant side effects were observed. Furthermore, Allon filed an IND for intravenously delivered NAP (AL-208). AL-208 is currently in phase II clinical trials. The objective of the trial was to secure safety data for the company to pursue CABG dementia as the lead indication for AL-208.

Acknowledgments Professor Illana Gozes is the incumbent of the Lily and Avraham Gildor Chair for the Investigation of Growth Factors. Supported by ISF, BSF, ISOA, NICHD, NIA, and Allon Therapeutics, Inc. NAP is under patent protection and licensed for clinical development to Allon Therapeutics, Inc. The intranasal formulation for NAP is termed AL‐108. Professor Illana Gozes serves as the Chief Scientific Officer of Allon Therapeutics, Inc. We would like to thank Drs. Bruce Morimoto, Anthony Fox, and Ms. Karole Sutherland for their help in the clinical development of AL‐108. Gozes I, Morimoto BH, Tiong J, Fox A, Sutherland K, Dangoor D, Holser-Cochav M, Vered K, Newton P, Aisen PS, Matsuoka Y, van Dyck CH, Thal L. 2005. NAP: Research and development of a peptide derived from activity-dependent neuroprotective protein (ADNP). CNS Drug Rev 11: 353-368.

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Gozes I, Littauer UZ. 1982. Microtubule protein: Tubulin. Scand J Immunol Suppl 9: 299-316. Gozes I, Alcalay R, Giladi E, Pinhasov A, Furman S, et al. 2002. NAP accelerates the performance of normal rats in the water maze. J Mol Neurosci 19: 167-170. Gozes I, Avidor R, Biegon A, Baldino F Jr. 1989. Lactation elevates vasoactive intestinal peptide messenger ribonucleic acid in rat suprachiasmatic nucleus. Endocrinology 124: 181-186. Gozes I, Bachar M, Bardea A, Davidson A, Rubinraut S, et al. 1997a. Protection against developmental retardation in apolipoprotein E‐deficient mice by a fatty neuropeptide: Implications for early treatment of Alzheimer’s disease. J Neurobiol 33: 329-342. Gozes I, Bachar M, Bardea A, Davidson A, Rubinraut S, et al. 1998. Protection against developmental deficiencies by a lipophilic VIP analogue. Neurochem Res 23: 689-693. Gozes I, Bardea A, Bechar M, Pearl O, Reshef A, et al. 1997b. Neuropeptides and neuronal survival: Neuroprotective strategy for Alzheimer’s disease. Ann N Y Acad Sci 814: 161-166. Gozes I, Bardea A, Reshef A, Zamostiano R, Zhukovsky S, et al. 1996a. Neuroprotective strategy for Alzheimer’s disease: Intranasal administration of a fatty neuropeptide. Proc Natl Acad Sci USA 93: 427-432. Gozes I, Davidson A, Gozes Y, Mascolo R, Barth R, et al. 1997c. Antiserum to activity‐dependent neurotrophic factor produces neuronal cell death in CNS cultures: Immunological and biological specificity. Brain Res Dev Brain Res 99: 167-175. Gozes I, Fridkin M, Hill JM, Brenneman DE. 1999. Pharmaceutical VIP: Prospects and problems. Curr Med Chem 6: 1019-1034. Gozes I, Giladi E, Pinhasov A, Bardea A, Brenneman DE. 2000a. Activity‐dependent neurotrophic factor: intranasal administration of femtomolar‐acting peptides improve performance in a water maze. J Pharmacol Exp Ther 293: 1091-1098. Gozes I, Glowa J, Brenneman DE, McCune SK, Lee E, et al. 1993. Learning and sexual deficiencies in transgenic mice carrying a chimeric vasoactive intestinal peptide gene. J Mol Neurosci 4: 185-193. Gozes I, Lilling G, Davidson A, Bardea A, Reshef A, et al. 1996b. Development of VIP agonists and antagonists with tissue and receptor specificity: Effects on behavioral maturation, sexual function, and the biologic clock. Ann N Y Acad Sci 805: 159-169; discussion 169-171. Gozes I, Lilling G, Glazer R, Ticher A, Ashkenazi IE, et al. 1995. Superactive lipophilic peptides discriminate multiple vasoactive intestinal peptide receptors. J Pharmacol Exp Ther 273: 161-167.

Gozes I, McCune SK, Jacobson L, Warren D, Moody TW, et al. 1991. An antagonist to vasoactive intestinal peptide affects cellular functions in the central nervous system. J Pharmacol Exp Ther 257: 959-966. Gozes I, Schachter P, Shani Y, Giladi E. 1988. Vasoactive intestinal peptide gene expression from embryos to aging rats. Neuroendocrinology 47: 27-31. Gozes I, Shani Y, Rostene WH. 1987. Developmental expression of the VIP gene in brain and intestine. Brain Res 388: 137-148. Gozes I, Vulih I, Spivak‐Pohis I, Furman S. 2005a. Neuroendocrine aspects of molecular chaperones: ADNF and ADNP. Henderson B, Packley AG, eds. Molecular chaperones and cell signalling. Cambridge University Press. pp. 251-262. Gozes I, Zaltzman R, Giladi E, Grigoriadis N, Brenneman D, et al. 2003. Treatment of experimental encephalomyelitis by intranasal application of ADNP peptide, NAP. Program No 9521. Abstract Viewer/Itinerary Planner Washington, DC: Society for Neuroscience. Gozes I, Zaltzman R, Hauser J, Brenneman D.E, Shohami E, et al. 2005b. The expression of activity‐dependent neuroprotective protein (ADNP) is regulated by brain damage and treatment of mice with the ADNP‐derived peptide, NAP, reduces the severity of traumatic head injury. Current Alzheimer’s Research. 2: 149-153. Gozes I, Zamostiano R, Pinhasov A, Bassan M, Giladi E, et al. 2000b. A novel VIP responsive gene. Activity‐dependent neuroprotective protein. Ann N Y Acad Sci 921: 115-118. Gressens P, Besse L, Robberecht P, Gozes I, Fridkin M, et al. 1999. Neuroprotection of the developing brain by systemic administration of vasoactive intestinal peptide derivatives. J Pharmacol Exp Ther 288: 1207-1213. Gressens P, Hill JM, Gozes I, Fridkin M, Brenneman DE. 1993. Growth factor function of vasoactive intestinal peptide in whole cultured mouse embryos. Nature 362: 155-158. Gressens P, Hill JM, Paindaveine B, Gozes I, Fridkin M, et al. 1994. Severe microcephaly induced by blockade of vasoactive intestinal peptide function in the primitive neuroepithelium of the mouse. J Clin Invest 94: 2020-2027. Gressens P, Marret S, Hill JM, Brenneman DE, Gozes I, et al. 1997. Vasoactive intestinal peptide prevents excitotoxic cell death in the murine developing brain. J Clin Invest 100: 390-397. Guo Q, Sebastian L, Sopher BL, Miller MW, Glazner GW, et al. 1999. Neurotrophic factors (activity‐dependent neurotrophic factor (ADNF) and basic fibroblast growth factor (bFGF)) interrupt excitotoxic neurodegenerative cascades promoted by a PS1 mutation. Proc Natl Acad Sci USA 96: 4125-4130. Hashimoto Y, Kaneko Y, Tsukamoto E, Frankowski H, Kouyama K, et al. 2004. Molecular characterization of

From concept to potential therapeutics: Neuroprotective peptides neurohybrid cell death induced by Alzheimer’s amyloid‐b peptides via p75NTR/PLAIDD. J Neurochem 90: 549-558. Ho AK, Chik CL, Klein DC. 1987. Transmembrane receptor cross‐talk: Concurrent VIP and a1‐adrenergic activation rapidly elevates pinealocyte cGMP greater than 100‐fold. Biochem Biophys Res Commun 146: 1478-1484. Ishihara T, Hong M, Zhang B, Nakagawa Y, Lee MK, et al. 1999. Age‐dependent emergence and progression of a tauopathy in transgenic mice overexpressing the shortest human tau isoform. Neuron 24: 751-762. Khan IU, Wallin R, Gupta RS, Kammer GM. 1998. Protein kinase A‐catalyzed phosphorylation of heat shock protein 60 chaperone regulates its attachment to histone 2B in the T lymphocyte plasma membrane. Proc Natl Acad Sci USA 95: 10425-10430. Lagreze WA, Pielen A, Steingart R, Schlunck G, Hofmann HD, et al. 2005. The peptides ADNF‐9 and NAP increase survival and neurite outgrowth of rat retinal ganglion cells in vitro. Invest Ophthalmol Vis Sci 46: 933-938. Lee VM, Goedert M, Trojanowski JQ. 2001. Neurodegenerative tauopathies. Annu Rev Neurosci 24: 1121-1159. Leker RR, Teichner A, Grigoriadis N, Ovadia H, Brenneman DE, et al. 2002. NAP, a femtomolar‐acting peptide, protects the brain against ischemic injury by reducing apoptotic death. Stroke 33: 1085-1092. Magistretti PJ, Schorderet M. 1984. VIP and noradrenaline act synergistically to increase cyclic AMP in cerebral cortex. Nature 308: 280-282. Maness LM, Kastin AJ, Weber JT, Banks WA, Beckman BS, et al. 1994. The neurotrophins and their receptors: Structure, function, and neuropathology. Neurosci Biobehav Rev 18: 143-159. Michaelis ML, Ansar S, Chen Y, Reiff ER, Seyb KI, et al. 2004. b‐Amyloid‐induced neurodegeneration and protection by structurally diverse microtubule‐stabilizing agents. J Pharmacol Exp Ther 48: 832-838. Morfini G, Pigino G, Beffert U, Busciglio J, Brady ST. 2002. Fast axonal transport misregulation and Alzheimer’s disease. Neuromolecular Med 2: 89-99. Offen D, Sherki Y, Melamed E, Fridkin M, Brenneman DE, et al. 2000. Vasoactive intestinal peptide (VIP) prevents neurotoxicity in neuronal cultures: Relevance to neuroprotection in Parkinson’s disease. Brain Res 854: 257-262. Ogiwara T, Murdoch G, Chik CL, Ho AK. 1995. Tyrosine kinase inhibitors enhance cGMP production in rat pinealocytes. Biochem Biophys Res Commun 207: 994-1002. Patterson PH. 1992. The emerging neuropoietic cytokine family: first CDF/LIF, CNTF, and IL‐6; next ONC, MGF, GCSF? Curr Opin Neurobiol 2: 94-97. Pinhasov A, Mandel S, Torchinsky A, Giladi E, Pittel Z, et al. 2003. Activity‐dependent neuroprotective protein: a novel

24

gene essential for brain formation. Brain Res Dev Brain Res 144: 83-90. Poggi SH, Goodwin K, Hill JM, Brenneman DE, Tendi E, et al. 2003. The role of activity‐dependent neuroprotective protein in a mouse model of fetal alcohol syndrome. Am J Obstet Gynecol 189: 790-793. Poggi SH, Vink J, Goodwin K, Hill JM, Brenneman DE, et al. 2002. Differential expression of embryonic and maternal activity‐dependent neuroprotective protein during mouse development. Am J Obstet Gynecol 187: 973-976. Romano J, Beni‐Adani L, Nissenbaum OL, Brenneman DE, Shohami E, et al. 2002. A single administration of the peptide NAP induces long‐term protective changes against the consequences of head injury: Gene atlas array analysis. J Mol Neurosci 18: 37-45. Said SI. 1991. Vasoactive intestinal polypeptide (VIP) in asthma. Ann N Y Acad Sci 629: 305-318. Said SI, Mutt V. 1970. Polypeptide with broad biological activity: I solation from small intestine. Science 169: 1217-1218. Schaad NC, Vanecek J, Rodriguez IR, Klein DC, Holtzclaw L, et al. 1995. Vasoactive intestinal peptide elevates pinealocyte intracellular calcium concentrations by enhancing influx: Evidence for involvement of a cyclic GMP‐dependent mechanism. Mol Pharmacol 47: 923-933. Sigalov E, Fridkin M, Brenneman DE, Gozes I. 2000. VIP‐ related protection against iodoacetate toxicity in pheochromocytoma (PC12) cells: a model for ischemic/hypoxic injury. J Mol Neurosci 15: 147-154. Sigurdsson EM, Lee JM, Dong XW, Hejna MJ, Lorens SA. 1997. Bilateral injections of amyloid‐b25–35 into the amygdala of young Fischer rats: Behavioral, neurochemical, and time‐dependent histopathological effects. Neurobiol Aging 18: 591-608. Smith‐Swintosky VL, Gozes I, Brenneman DE, D’Andrea MR, Plata‐Salaman CR. 2005. Activity‐dependent neurotrophic factor‐9 and NAP promote neurite outgrowth in rat hippocampal and cortical cultures. J Mol Neurosci 25: 225-238. Spong CY, Abebe DT, Gozes I, Brenneman DE, Hill JM. 2001. Prevention of fetal demise and growth restriction in a mouse model of fetal alcohol syndrome. J Pharmacol Exp Ther 297: 774-779. Steingart RA, Solomon B, Brenneman DE, Fridkin M, Gozes I. 2000. VIP and peptides related to activity‐dependent neurotrophic factor protect PC12 cells against oxidative stress. J Mol Neurosci 15: 137-145. Takahashi RH, Almeida CG, Kearney PF, Yu F, Lin MT, et al. 2004. Oligomerization of Alzheimer’s b‐amyloid within processes and synapses of cultured neurons and brain. J Neurosci 24: 3592-3599.

687

688

24


White DM, Walker S, Brenneman DE, Gozes I. 2000. CREB contributes to the increased neurite outgrowth of sensory neurons induced by vasoactive intestinal polypeptide and activity‐dependent neurotrophic factor. Brain Res 868: 31-38. Wilkemeyer MF, Chen SY, Menkari CE, Brenneman DE, Sulik KK, et al. 2003. Differential effects of ethanol antagonism and neuroprotection in peptide fragment NAPVSIPQ prevention of ethanol‐induced developmental toxicity. Proc Natl Acad Sci USA 100: 8543-8548. Wilkemeyer, MF, Chen, SY, Menkari, CE, Sulik, KK, Charness, ME. 2004. Ethanol antagonist peptides: Structural specificity without stereospecificity. J Pharmacol Exp Ther 309: 1183-1189. Wilkemeyer MF, Menkari CE, Spong CY, Charness ME. 2002. Peptide antagonists of ethanol inhibition of L1‐mediated cell–cell adhesion. J Pharmacol Exp Ther 303: 110-116. Wu JY, Henins KA, Gressens P, Gozes I, Fridkin M, et al. 1997. Neurobehavioral development of neonatal mice following blockade of VIP during the early embryonic period. Peptides 18: 1131-1137. Zaltzman R, Alexandrovich A, Beni SM, Trembovler V, Shohami E, et al. 2004. Brain injury‐dependent expression of activity‐dependent neuroprotective protein. J Mol Neurosci 24: 181-187.

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Index

A disintegrin and metalloproteinase (ADAM), 566–567, 570–574, 577–580 A disintegrin and metalloproteinase with thrombospondin repeats (ADAMTS), 567, 569–573, 577 Acetyltransferase coenzyme A2, 275 Acetyltransferase Tip60, 275 aCGH, 273–274 Actin, 223 – ADP-bound form, 226, 230 – ATP-bound form, 226, 227, 230 – filamentous (F-), 225 – globular (G-), 225, 228 – g-isoform, 225 – polarity of, 226–228 – polymerization of, 226, 238 Actin filament (F-actin) – barbed end of, 228, 230, 231 – double helix of, 225 – pointed end of, 228, 237 – treadmiling of, 225 Actin polymerization, 228, 230–232, 238 – critical concentration of, 225, 226 – elongation phase of, 226, 227 – equilibration phase of – nucleation phase of, 226, 227 a-actinin, 234, 238 – EF-hand structure of, 233 – F-actin crosslinking activity of, 231, 233 Activity-dependent neuroprotective protein (ADNP), 674–677, 680, 681 Activity-dependent neurotrophic factor, 674 – ADNF‐14, 675, 676, 678 – ADNF‐9, 675–682, 684 – SAL, 676–684 Adenylyl cyclase, 270 ADF/Cofilin, 229

#


Adhesion molecules, 285, 286, 290 – L-selectin, 288, 291–294, 297 – peripheral node vascular addressin (PNAd) 284, 291–293 – P-selectin, 284, 291, 294 – P-selectin glycoprotein ligand 1 (PSGL‐1), 284, 291, 293, 294 – sulfoadhesin, 294 Adseverin, 228 Advillin, 228 Aggrecanase, 570 Aggregates, 659 Aggregation, 356, 357, 360–364, 366 Aging, 659 Aging pigment, 666 Atg7 mice, 668 ALCAM, 4, 80–83, 108, 111 Allan-Herndon-Dudley syndrome, 267 Alzheimer’s disease (AD), 214, 347, 349, 447, 448, 470, 473, 578, 579, 592, 593, 601, 604, 606, 658, 659, 674, 681–684 – amyloid peptides, 454, 675, 678, 679, 682 – calpain proteolytic system, 455 – conversion of p35 to p25, 455 – dementia, 454 – hippocampal neurons, 454–456 – plaques and tangles, 454 – sporadic AD (SAD) or familial AD (FAD), 595, 605 AMICA, 41, 53, 55–57 Aminopeptidase, 626, 629–640, 642, 643 Ammon’s horn sclerosis (AHS), 198, 210, 213 Amyloid, 578–579 Amyloid b (Ab), 578, 592 – Ab degradation, 594, 596, 598, 605, 606 – Ab fibrillation, 596 Amyloid peptide, 659

Amyloid peptide, Ab, 470 Amyloid precursor protein (APP), 470, 473–484, 486–493, 495–497, 570 – a-secretase, 578 Amyloid precursor, 275 Amyloidosis, 327 Amyotrophic lateral sclerosis (ALS), 195, 214, 448 – glutamate toxicity, 455 – Lou Gehrig’s disease, 455 – motoneuron degeneration, 455 – oxidative stress and Ca2+ influx, 455 Anaphase-promoting complex, 656 Angelman’s syndrome, 266 Angiogenesis, 295, 576, 577 – angiogenic switch, 294, 296 Angiotensin II-specific, 269 Angiotensin, 626, 629, 633, 634, 641, 643 APC, 656 Apolipoprotein E, 330, 596, 629, 678 Apoptosis, 658, 659 – death receptor-mediated pathway, 518, 519, 525, 526 – endoplasmic reticulum-mediated pathway, 526–528 – mitochondrial pathway, 521, 522, 529 – morphological and biochemical features, 516, 517, 519, 520, 530 – role of caspases, 530 Apoptosis-inducing factor (AIF), 517 – participant in DNA degradation, 520, 521 – release from mitochondria in apoptosis, 520, 530 – role in caspase-independent apoptosis, 530 Apoptosome, 520

690

Index Apoptotic protease activating factor (Apaf)‐1, 517 – role in caspase‐9 activation, 520, 521 Arginine vasopressin, 270 Argininosuccinase, 264 Aristaless-related homeobox, 266 Arp2/3 complex – F-actin branching activity of, 232, 238 – increase of polymerization point, 231, 232, 238 – N-WASP binding of, 231, 232, 238 Array comparative genomic hybridization, See aCGH ARX protein, 266 Aspartic proteases inhibitors – pepstatins, 606 Atg5, 668 Atg7 mice, 668 Autism, 261, 271, 272, 276 Autofluorescence – as property of lipofuscin, 666, 667 Autophagosomes, 665 – role in garbage catastrophe, 664, 666–668 Axon – elongation, 657 – guidance, 657 – initiation, 657 – regeneration, 656 Ab peptides, 552–554, 558 – intracellular degradation of, 555, 556 – extracellular degradation of, 559 BAPTA, 200–203, 205, 210 Bcl‐2 family, 529 – Bcl‐2 homology (BH) domains, 522, 523 – member subclasses, 522, 523 – overexpression as potential therapeutic agent, 534 – regulation of apoptosis, 521–523 Bestatin, 633, 636, 637, 640, 642–644 Biogenesis, 154, 155, 161, 162 – dense-core secretory granules, 163, 164 Bipolar disorder, 212 Blood-brain barrier, 574–576, 578–580 Brain peptidase, 626 Brain tumors, 592, 602, 603, 605 Brain-derived neurotrophic factor, 158 – activity-dependent secretion of, 154

– carboxypeptidase E, sorting receptor for, 157 – sorting signal in, 155–157 Branched-chain a-keto acid dehydrogenase (BCKAD), 263, 264, 276 Ca2+ binding proteins, 447 Ca2+ extrusion and sequestration, 448 – Ca2+-ATPase, 449, 450 – energy depletion, 450 – mitochondrial sequestration of Ca2+, 450 Calbindin D‐28k (CB), 198, 205–208, 451, 456 Calcium – binding protein, 198–215 – buffer, 200–205 – sensor (modulator), 198–200, 204 – signaling, 198, 204, 208, 210, 215 Caldesmon – calmodulin binding of, 234 – inhibit actin polymerization promoting activity of N-WASP, 231, 232, 238 – tropomyosin binding of, 234 Calmodulin (CaM), 198, 199, 205, 451 Calpain, 559, 560, 657 – cerebral ischemia, 20 Calpain activation, 457, 459 – failure of energy-dependent mechanisms, 458 – inhibition, 451–456, 458 – intraneuronal Ca2+ overload, 449, 451 – multiple mechanisms, 447–449, 458 Calretinin (CR), 198, 199, 205, 276 CAM, 56, 59–61, 63, 66, 69, 72, 77, 79, 80, 85, 86, 88, 90, 97, 99, 101, 103, 105–108, 111–113, 115, 117–119 CapG, 228 Carboxypeptidase E, 158, 159 – association with lipid rafts, 157, 160, 162 – amphipathic a-helix, 157, 160 – CPEfat/fat mice, 157 – knock-out mouse, 157 – sorting receptor for neuropeptides and BDNF, 157 – transmembrane protein, 157, 160 Carboxypeptidase, 626, 628–632, 634, 641 Caspases, 560, 561 – activation in cell death, 517, 519 – cellular substrates, 529, 533

– competitive inhibitors as potential therapeutic agents, 533, 534 – domain structure, 517, 518 – family subclasses, 517–519 – non-apoptotic functions, 530 Caspr, 41, 102, 103, 112 Catalytic b-subunits, 657 Cathepsins, 553–556 – alterations in aging, 666 – role in lysosomal degradation, 664, 665, 667 c-Cbl, 656, 657 cCGH, 273 CD47, 49–53, 63–66 cDNA array, 274 Cell adhesion L1-like, 266, 272 Cell adhesion molecule (CAM), 41–43, 46, 55, 56, 59–61, 63, 66, 69, 72, 77–80, 85, 86, 88–90, 96, 97, 99, 101, 103, 105–108, 111–115, 117–120 Cell death, 658, 659 Centenarians, proteolysis in, 666 Central nervous system (CNS) trauma – behavioral recovery, 534, 536 – brain injury, caspase activation, 531, 533 – ischemic brain injury (stroke), 531 – neuroprotection, 530, 535, 536 – rodent models, 531–533 – spinal cord injury, 532, 543 – therapeutic strategies, 533, 534 Central nervous system (CNS), 198–200, 205, 206, 212, 215, 296 – axonal guidance, 298 – injury, 297, 298 CEPU‐1, 41, 79 Cerebral amyloid angiopathy (CAA), 592, 593, 595, 596, 601 Cerebral infarction, 597, 601 – cerebrovascular deposits (CD), 470 Chaperone, 270, 378, 384, 388, 392, 394, 396, 654 – heat shock stress, 390 Chemokine receptors, 284, 290, 291, 293 Chemokines, 285, 295, 566–568, 573, 574, 580 CHL1, 41, 105–108, 110, 112, 113 Cholesterol – cholesterol-sphingolipid-rich membrane microdomain, 157 – dense-core secretory granules, 160, 162–164 – lathosterolosis mouse, 161, 162 – lipid rafts, 157, 160, 162, 164 – Smith-Lemli-Opitz syndrome, 161

Index Chromogranin A – assembly factor, 154, 162 – controlling stability of granule proteins, 163 – degradation of granule proteins, in the absence of, 163, 164 – depletion of, 161, 163 – granulogenic role of, 162, 163 – regulation of dense-core secretory granule biogenesis by, 163, 164 Chronic inflammation, 291–293 cis-interactions, 44, 47, 49, 51, 69, 70, 75, 90, 91, 117 CLMP, 41, 53 CLN1, 664, 665 – mutation in NCL, 667 Collagenase, 568 Comparative genomic hybridization, See cCGH Conformational change, 654 Congenital disorders of glycosylation, 264, 265 Congo red, 396 Contactin, 101–105, 114 Convertase, 626, 631, 632 Converting enzyme, 626, 629 20S core, 655 Creatine defects, 262, 263 Creatine kinase, 276 CREB, 268 CRE-binding, 268 Creutzfeldt-Jakob disease, 601 Crinophagy, 665 CTX, 41, 53–55, 58, 70 CXADR, 41, 53–58 Cyclin D1, 659 Cyclin dependent kinase-like, 271 Cyclosporin A, 535 Cysteine proteases inhibitors, cystatins, 601 – kininogens, 599, 600 – stefins, 599, 603 Cystic fibrosis transmembrane conductance regulator, 391 Cytochrome b5, 276 Cytochrome c, 517, 523, 526–528, 531, 532, 534, 536 – release from mitochondria in apoptosis, 520–522 – role in caspase‐9 activation, 520, 521 Cytoskeleton, 176, 178 Cytotoxic proteins, 659 Dasm1, 119 DCC, 41, 99, 100, 120 Death-inducing signaling complex (DISC), 526

Death receptors, 526 – apoptotic pathway, 521, 522, 529, 530 – caspase activation, 528 Degenerative disease, 626 Degradation products, 655 3p Deletion syndrome, 272 Dendrite, 170–176, 178–188 Dendritic cells, 293 Dendritic growth, 656 Dendritic spine, 170 Dense-core secretory granule – aggregation of cargo proteins in, 162, 163 – biogenesis of, 163 Deubiquitinating enzymes, 654, 655, 669 – association with neurodegeneration, 668 Dihydropteridine reductase, 262 Dipeptide-inhibitor, 656 Disaggregation, 658 DJ‐1, 658 Down syndrome, 261, 275, 592 Drebrin – inhibit fascin activity, 232 – inhibit myosin activity, 234, 235–238, 255 – inhibit tropomyosin binding to F-actin, 234 DSCAM, 41, 113–117 DSCAML1, 41, 113–116 DUB, 655, 656 E6-AP, 658, 659 4E-BP1, 10, 12, 16, 19, 24 E-cadherin, 271 EF-hand, 198–201, 215 EGFR, 42, 64 EGTA, 205, 210 eIF2, 4–6, 9, 13, 14, 16–21, 24 eIF2B, 9, 12, 16, 17, 19–21 eIF4G, 4–7, 10, 11, 13, 14, 16, 17, 19–21, 23 Endocytosis, 654–657 – age-associated change in, 665 Endonuclease G (Endo G), 517 – participant in DNA degradation, 520, 521 – release from mitochondria in apoptosis, 520, 521 – role in caspase-independent apoptosis, 530 Endopeptidase, 626, 629, 630–632, 634 Endoplasmic reticulum (ER), 174, 346, 519 – apoptotic pathway, 526–528 – cross-talk with mitochondria, 528

– endoplasmic reticulum stress, 347, 351 – unfolded protein response (UPR), 347, 348, 351, 352, 526 Endoplasmic reticulum (ER) stress, 9, 14, 356, 359, 362, 658 Endosomal membrane, 656 Endothelial cells, 291–293, 295, 296 Energy landscape, 308–310, 313–315, 318 Enkephalin, 626–628, 630–636, 638, 641–644 Epilepsy, 210, 212 Epileptic seizure, 448, 597 – abnormalities in electrophysiological activity, 456 – hyperexcitability, 456 – oxidative stress, 445, 456–458 – oxygen-regulated protein‐150 kDa, 447, 456 – seizure-induced brain damage, 456 Endoplasmic reticulum associated degradation (ERAD) – connection to cytoplasmic degradation, 667 ESAM, 42, 53–55, 58, 133 Excitotoxicity, 213, 214 Executioner caspases, 518–520, 526, 527, 529 Exogenous antigen presentation – class II-associated Ii peptide (CLIP), 552, 553 – invariant chain (Ii), 453, 460 – major histocompatibility complex (MHC) class II, 552, 553 – ovalbumin (OVA), 553, 554 – T helper cell hybridomas, 553 Exopeptidase, 629, 630, 641 Experimental allergic encephalitis (EAE), 575 Extracellular matrix – collagen, 566–570, 572, 580 – laminin, 569 – proteins, 553–555 – proteoglycans, 566, 568–570, 572 F3, 41, 44, 61, 101–104, 107, 108, 111, 112 Familial AD (FAD), 470 Familial encephalopathy, 594, 597 Fascin – C-kinase regulation of, 232 – F-actin bundling activity of, 231, 232 – formation of polarized F-actin bundle, 232 Fetal alcohol syndrome, 678, 680

691

692

Index Fetoprotein, 262 FGFR, 42, 79, 91, 92, 94, 107–109, 134 Fibronectin, 569, 579 FISH, 272, 274 FLRT, 42 Fluorescence in situ hybridization, See FISH FMR1, 265 FMRP, 265 Fragile X mental retardation protein (FMRP), 175, 176, 185 Fragile X syndrome, 265, 277 Fucosylated, 265 Fukutin, 272 Fukuyama congenital muscular – dystrophy, 271, 272 G proteins, 267, 270 GABA receptor Ag2, See GABRG2 GABRG2, 275 GADD34 expression, 19, 24 Galactokinase, 263 Galactose‐1-phosphate uridyl transferase, 263 Galactosemia, 263 Garbage catastrophe, 664, 668 – proposed mechanism, 666, 667 GDP dissociation inhibitors, 267 GDP-fucose transporter, 265 Gelatinase, 569, 580 – gelatinase A, 568, 575 – gelatinase B, 568 Gelsolin – calcium regulation of, 228 – F-actin capping activity of, 228, 229, 230, 234 – F-actin severing activity of, 228, 229 – nucleation promoting activity of, 228, 229 – PIP2 regulation of, 228, 230 Gene therapy, 536 GGEL, 244, 246, 247, 251–253 Glia, 566, 568, 575 – astrocytes, 569, 572, 573, 576–578, 580, 581 – microglia, 569, 572–574, 579 – oligodendrocytes, 576 Glucocorticoids, 568 Glutamate receptors – a-Amino‐3-hydroxy‐5-methyl‐ 4-isoxazole propionate receptor, 447, 448 – Ca2+ influx, 448–450, 457 – endoplasmic reticulum, 448, 449 – ionotropic, 447, 448, 455, 457 – metabotropic, 447–449, 455 – N-methyl-D-aspartate receptor, 447, 448

Glycolipids, 284, 286, 290, 298 – sulfatide, 296, 297 Glycoproteins, 264, 284–286, 290, 291, 296, 298 – mucins, 293 Golgi apparatus, 174, 189 Granulogenic proteins, 162 Growth cone dynamics, 657 Growth cone, 225, 226, 232 – C-domain – dynamics of, 236, 238 – filopodium, 228, 229, 231, 236–238 – lamellipodium, 228, 229, 231, 237–238 – P-domain, 236 Growth factors, 284, 294–296 GRP78, 276 GSTO1, 276 GTPase, 267, 268 GTP-binding proteins, 267 Guanidinoacetate methyltransferase, 262 HADII, 263 HDAC inhibitor, 265 HECT domains, 654 Helicase 2, 267 Hemoglobin, 266 Hereditary cerebral hemorrhage with amyloidosis (HCHWA), 595 – Dutch type (HCHWA-D), 592 – Icelandic type (HCHWA-I), 601 Heterophilic interaction, 44, 59, 79, 107, 113, 118, 120 High endothelial venules (HEV), 284, 291–292, 294 – extralymphoid HEV-like vessels, 293 Hippocampus – dentate gyrus, 569, 571 Histone acetyltransferase, 268, 275 Histones, 276 HLA-DRB3, 275 Holoprosencephaly, 269 Homophilic interactions, 44, 47, 50, 56, 58, 60, 63, 69, 82, 85, 87, 90, 96, 99, 102, 104, 107, 111–113, 115, 118 Hormone half-life, 290, 296 – lutropin, 284, 295 – thyrotropin, 284, 295 HR6, 656 Hsc73 – role in lysosomal degradation, 665 HtrA2/Omi, 517, 530 – endogenous inhibitor of IAPs, 520 – release from mitochondria in apoptosis, 520 Hunter syndrome, 264

Huntingtin, 659 Huntington’s disease, 448, 668 – CAG repeat, 457 – chorea – Huntingtin, 457 – neurodegeneration, 457 – polyglutamine disease, 457 – striatum, 457 Hybrid proteasome, 378, 379 Hydroxysteroid dehydrogenase, 263 Hypoparathyroidism-retardationdysmorphism syndrome, 270 Iduronate sulfatase, 264 IgLON, 42, 45, 77–80 Ig-module, 44–51, 53–57, 60–63, 68, 70, 74, 76, 79, 82–84, 87, 89, 90, 92, 96, 98, 99, 101–102, 104, 107, 109, 112, 114–116, 118–120 IGSF, 41–47, 75, 77, 120 IGSF11, 53, 55, 56, 58, 59, 133, 149 IGSF9, 42, 118, 119 IL1RAP, 269 Immediate early gene (IEG), 170, 178 Immunoglobulin superfamily, 42–44, 46, 55, 73, 119 Immunoproteasome – LMP2, 376, 399 – LMP7, 376, 399 – LMP10, 376 Inclusions, 659 Inflammation, 568, 569, 572, 573, 578–580 Inhibitor, 626, 629, 630, 633–644 Inhibitors of apoptosis (IAP), 518, 528 – endogenous inhibitor of caspases, 520 – overexpression as potential therapeutic agent, 534, 535 Initiation factors, 3–6, 8, 10, 11, 16, 17, 19 Initiator caspases, 518, 519, 526, 528 Insoluble aggregates, 659 Insulin, 270 Insulin-degrading enzyme, 559 7-Integrin, 272 Integrin-associated protein, 49–53 Interferon-b, 574 Interleukin‐1 receptor accessory, 269 Interleukin‐1b, 573 Internalization, 657 Intracellular inclusions – aggresomes, 373, 391, 396, 397 – centrosome, 387, 391, 392, 397 – degrasome, 396 – g-tubulin , 387, 390–392 – microtubules, 388, 391, 397 – neuronal intanuclear inclusions, 373, 384

Index – proteolysis center, 391, 396 – tubulin, 387, 388, 390–392 Intracerebroventricular (ICV) injections, 533, 534 Ischemic brain injury – blockage of cerebral blood supply, 451 – calpain inhibitor MDL‐28170, 452 – calpain-mediated proteolysis, 452 – excitotoxicity, 451 – ischemic penumbra, 451, 452 – necrosis and apoptosis, 451 Isovaleryl-CoA dehydrogenase, 276 Kilon, 42, 77–79 L1, 41, 42, 47, 75, 85, 87, 92, 103–113 L1CAM, 266, 272 Lactacystin, 658 Lamin, 276 Lamp2a, role in chaperone-mediated proteolysis, 665 LC-MS/MS, 276–277 Leukocyte recirculation, 291, 297 Lewy bodies, 659 Ligand-gated Ca2+ channels, 450 – Ca2+ releasing channels, 449 – IP3 receptors, 448, 449 – ryanodine receptors, 447, 449 Ligase E3A, 266 Lipid rafts, carboxypeptidase E, 158, 159 – cholesterol-sphingolipid-rich microdomain, 157 Lipofuscin – association with aging, 666 – role in garbage catastrophe, 666, 667 Lipopolysaccharide, 573 Lissencephaly‐1, 264 LMP2 – mouse knockout model, 668 Long term depression (LTD), 170 Long-term potentiation (LTP), 170, 171, 178, 182, 185–186 Lymphoid organs – lymph nodes, 293 – peripheral lymphoid organs, 291 Lysosomes, 654, 664, 665, 667 Macroautophagy, 665 a‐macroglobulins, 572, 593 MAG, 41, 47, 56, 86–88 Maintenance, 654, 656, 657, 659 MALDI-TOF-MS, 275 Malonyl-, Methylmalonyl-CoA – decarboxylase defects, 263 Malonyl-CoA decarboxylase, 263 MAPH, 274 Maple syrup urine disease, 263, 264, 276 MASA syndrome, 266

Matrilysin, 569 Matrix assisted laser desorption/ ionization time-of-flight – mass spectrometry, See MALDI-TOF – MS, 275, 276 Matrix metalloproteinase (MMP), 558, 559, 567, 570 – collagenase, 568 – gelatinase, 568, 569 – matrilysin, 569 – membrane-type MMP, 566–572 – other MMPs, 569, 570 – stromelysin, 569 mDia1, actin polymerization promoting activity of, 231 MECP2, 271, 276 Melanoma cell adhesion molecule (MCAM), 41, 42, 83–86 Metalloproteases inhibitors, 606 – tissue inhibitors of metalloproteases (TIMP), 592, 604, 605 Methyl CpG-binding, 271 Methylmalonyl-CoA decarboxylase, 263 MHC DR b3, See HLA-DRB3 Microdeletions, 268, 273, 274 Microduplications, 273, 274 Microglia, 552–556, 559, 560, 568, 569, 572–574, 579 – activation of, 557, 558 – apoptosis of, 561 Microtubule, 659, 677, 682, 683 Misfolding, 659 Mitochondria, 356, 362, 363, 366, 535, 665 – apoptotic pathway, 520–524 – caspase activation, 528, 529 – contribution to lipofuscin, 666 – degradation by autophagy, 668 – source of reactive oxygen, 667 – cross-talk with endoplasmic reticulum, 522 – cross-talk with nucleus, 520 – dysfunction in apoptosis, 526 – inhibition of permeability transition as potential therapy, 523, 524 – release of pro-apoptotic factors, 520–523 Molecular dynamics, 312, 313 Molten globule, 310–332 Monocarboxylate transporter‐8, 267 Monosomy, 273 Mono-ubiquitination, 654, 656, 657

Mov34 mice, loss of function in proteasome, 668 Mowat-Wilson syndrome, 271 mRNA, 170–176, 178, 180–189 – structural elements, 6, 7 mTOR, 3, 10–12 Multiple sclerosis, 448, 593, 598, 600, 606 – autoimmune inflammatory reaction, 458 – demyelination, 458, 459 – destruction of myelin proteins, 458, 459 – experimental allergic encephalomyelitis – neurodegenerative component, 447, 459 – white matter plaque, 458 Multiplex amplifiable probe hybridization, See MAPH Multi-ubiquitination, 654 Mutated proteins, 654 Myelin protein zero, 41, 45–49 Myelin-associated glycoprotein, 41, 42, 86–88 Myosin – -I, 236 – -II, 235–237 – -III, 237 – -V, 236 – -VI, 237 – Snell’s waltzer deafness, 237 – -VII, 237 – Usher 1B syndrome, 237 – -IX, 237 – -X, 237 – ATPase activity of, 234–236 – coiled-coil structure, 235, 237 – head domain, 235, 237 – heavy chain, 235, 237, 238 – IQ motif, 236, 237 – light chain phosphorylation, 235, 236 – neck domain, 235, 237 – phosphorylation by myosin light chain kinase, 235 – phosphorylation by rho kinase, 235 – processive movement, 237 – tail domain, 236, 237 NAP, 676–684 NCAM2, 42, 95–97 N E-(g-L-glutamyl)-L-lysine, 244, 246, 247 Necl, 42, 45, 66, 67, 72–77 Nectin, 45, 66–72, 74–76 Nectin-like, 71–77 Nedd4, 657 N-end rule, 656

693

694

Index Netrin G1, 271 Neural cell adhesion molecule (NCAM), 41–43, 47, 57, 63, 65, 75, 79, 85, 87–95, 108, 109, 119 Neurite outgrowth, 656, 657 Neurodegeneration, 249, 253, 254, 356, 363, 366, 659 Neurodegenerative disease – b-amyloid in, 346 – polyglutamine disease, 349, 350 – prion disease, 350 – synuclein in, 349, 350 – tau in, 346–349 Neurodegenerative, 214 Neurofascin, 42, 102–108, 110–112 Neurological diseases, 566 – Alzheimer’s disease, 570, 578, 579 – amyotrophic lateral sclerosis, 581 – glioma, 577, 581 – anaplastic astrocytomas, 576 – glioblastoma multiforme, 576 – human immunodeficiency virus dementia (HIVD), 579, 580 – multiple sclerosis, 572, 574, 575 – Parkinson’s disease, 571 – polyglutamine disease, 581 – stroke (cerebral ischemia), 575, 576 Neuron, 200, 201, 204–212, 214, 215, 654–659 Neuronal calcium sensor (NCS), 199 Neuronal ceroid lipofuscinoses, 667 Neuronal development, 655–657 Neuronal survival, 659 Neuropeptidase, 626, 644 Neuropeptides, 161, 626–638, 640 – regulated secretion of, 157–159 – sorting signals in, 154–157, 164 Neuroplastin, 59, 60 Neurotoxicity, 578, 579 NMDA receptor complex (NRC), 170, 176 NMDA receptors, 557 Nope, 42, 119, 120 NrCAM, 42, 82, 83, 102–108, 110–112, 119 N-WASP – actin polymerization promoting activity of, 231, 232, 238 – Arp2/3 binding of, 231, 232, 238 – Cdc42 regulation of, 231 OBCAM, 42, 77–80 OCAM, 42, 95 Oligonucleotide arrays, 274, 275, 277 Oligophrenin, 267 Opioid peptide, 630, 634, 644 Ornithine transcarbamylase, 264 Oxidative damage, 659

P0, 45, 47–49, 63, 78, 85, 87 p21-activated kinase, 268, 269 P2X7 receptors, 560, 561 PA200, 379 – ARM/HEAT repeats, 377 PAK proteins, 268 Palmitoyl protein thioesterase‐1, mutation in NCL model, 667–669 Parkin, 658 Parkinson’s disease, 447, 448, 596, 658, 659 – degeneration of dopaminergic neurons, 457 – Lewy bodies, 458 – movement disorder, 457 – substantia nigra, 457, 458 – a-synuclein, 458 Partington syndrome, 266 Parvalbumin (PV), 198, 199, 205–215 Patched, 269 PDGFR, 42, 64, 76 PDZ, 42, 56–58, 67, 68, 70, 74, 76, 111, 118, 119 Peptide – degradation, 374 – processing, 632 Peroxisomes, 665 PGP9.5, 659 Phenylalanine hydroxylase, 262 Phenylketonuria, 261 Phosphoadenosine‐5´-phosphosulfate (PAPS), 284, 286 – PAPS synthase, 287 Phosphomannomutase, 261 Phosphorylation, 654, 657 PI31, 658 PKU, See Phenylketonuria Plasmin, 556–558, 560 Plasminogen activators, 556–558, 560 Platelet-activating factor acetylhydrolase, 264 Poliovirus receptor-related, 42, 66, 68 Polyamines, 244, 249, 252 Polyglutamine expansion diseases, 391 – ataxin‐1, 393, 395, 397, 398 – ataxin‐2, 394 – CAG repeat, 393, 394 – dentatorubral pallidoluysian atrophy, 393–394 – Huntingtin, 381, 385, 386, 393–399 – Huntington’s disease, 373, 380, 386, 390, 393, 396, 397, 399 – Machado-Joseph disease, 393 – polyQ tract, 393–399 – spinobulbar muscular atrophy (Kennedy’s disease), 393 – spinocerebellar ataxias, 373, 393

Polyglutamine, 357–366 – binding, 265, 266 Polypyrimidine-tract binding protein – dense-core secretory granule biogenesis, 161–164 – post-transcriptional mechanism, 164 – stability of mRNAs encoding granule proteins, 164 Polyribosomes, 170–174, 180, 186, 188 Poly-ubiquitination, 654 Postsynaptic density, 180, 182 Prader-Willi syndrome, 265, 266 Presenillin, 391 Procaspase, 517, 519, 520, 526, 528, 529, 535 Process outgrowth, 659 Procollagen N-proteinases, 567, 570 Profilin, 226, 229 – ATP/ADP exchange, 230 – sequestering activity of, 230 Progressive myoclonus epilepsies (PME), 603 Prohormone convertase 1/3, 164 – amphipathic a-helix in, 160 – association with lipid rafts, 160 – C-terminus of, 160 – cytoplasmic tail of, 157, 158 – pro-domain of, 160 – sorting information in, 160 – trafficking of, 157 – transmembrane protein, 160 Prohormone protease, 632 Proinsulin – carboxypeptidase E, 157, 158 – hexamerization of, 155 – sorting signal in, 155–157 – sorting to regulated secretory pathway, 154, 156, 158, 159 – X-ray crystal structure of, insulin, 155, 156 Proopiomelanocortin, 154 – carboxypeptidase E, 157–159 – NMR structure for, 155 – sorting signal in, 155, 157 Pro-protein convertase, furin and furin-like proteases, 568, 571 Protease catalytic mechanism, 410–413 Protease classification of proteases, families and clans, 410, 411 Protease, human trypsin 4, 411, 430, 434, 435 – biochemical characterisation of 4, 431–433 – peptide substrates, 433 – pH-jump activation, 432 – polypeptide substrate, 433

Index – reaction with MUGB, 432 – resistance to canonical inhibitors, 433 – restricted substrate specificity, 413, 433 – small synthetic substrates, 432, 435 – chromosomal localization, 414, 428, 429 – expression in different areas of brain, 429–431 – immunohistochemical localization, 431 – isoforms of human trypsin 4, 428, 429 – alternative splicing, 419–425 – alternative transcription initiation, 429 – CUG start codon, 428, 429 – leader peptide, 428–431 – leucine as the initiator amino acid, 429 – 5´RACE PCR, 429 – regulation of expression level, 417, 419, 429 – potential roles in the CNS, 426, 427 – Alzheimer’s disease, 410, 418, 427 – cancer, 413, 425, 426 – cell migration, 413, 416 – metastasis, 426, 434 – migration of GnRH neurons, 434 – multiple sclerosis, 410, 426 – myelin degradation, 427 – PAR activation, 433 – tumour suppression, 427 – structure of human trypsin 4, 411, 412, 432 – activation peptide, 418, 425 – Arg193 (S2’ subsite), 432 – disulfide bridges, 412, 413 – leader peptide, 428,–431 – oxyanion hole, 411, 432, 433 – substrate binding site, 412 Protease, kallikreins, 420 – alternative splicing, 419, 425 – association of kallikreins with diseases – cancer, 425, 426 – hypertension, 426 – pathological keratinization, psoriasis, 426, 427 – renal nephritis, 426 – serum tumor biomarkers, 425, 426

– biological roles of kallikreins, 426 – activation of uPA and plasminogen, 410, 413 – inflammation, 413, 426, 433 – ischemia, 425, 427 – processing biologically active peptides, 425 – regulation of blood pressure, 425 – chromosomal localization, 414, 428, 429 – classification, plasma and tissue kallikreins, 410 – expression in central nervous system, 428 – gene structure of kallikreins, 418 – potential roles of kallikreins in the central nervous system – Alzheimer’s disease, 427 – demyelination, 426, 433 – experimental autoimmune encephalomyelitis, 410, 426 – frontotemporal dementia, 410, 428 – glutamate-receptor mediated cytotoxic injury, 426 – hippocampal networking, 427 – hippocampal plasticity, 427 – long-term potentiation, 410, 416 – multiple sclerosis, 410, 426 – multiple system atrophy, 410, 426 – myelination, turnover of myelin protein, 426 – neural plasticity, 427 – neurite outgrowth, 416, 427, 435 – neuronal development, 427 – neuroprotection in ischemia, 427 – Parkinson’s disease, 410, 426, 427 – production of cerebrospinal fluid, 427 – regulation of seizures, 427 – synaptogenesis, 427, 435 – synucleinopathies, 426, 427 – regulation of catalytic activity, 411 – regulation of expression, steroid hormones, 419 – sequence homology among kallikreins, 418, 419, 426 – structure of kallikrein enzymes, 418 – activation peptide, 418, 425

– disulphide bridges, 412 – kallikrein loop, 425 – signal peptide, 411, 418, 425 – substrate binding site, 412 Protease activated receptors (PAR), 414, 425, 433, 434 – activation of PARs by proteases and, 416 – expression of PARs in central nervous system, 415, 416 – vinduction of cell death, 417, 418 Protease, plasma serine – activity and specificity of haemostatic serine proteases, 413, 414 – blood clotting, 413 – brain damage, 417 – chromosomal localization of haemostatic serine proteases, 414 – coagulation cascade, 413, 414 – role of plasma serine proteases in the CNS, 417–418 – development, 413–416 – neuronal survival, 416 – plasticity, 416 – thrombin – Alzheimer’s disease, 418, 427, 428 – expression in different tissues, 415, 416 – facilitation of long-term potentiation, 416–418 – induction of apoptosis, 418 – induction of edema, 418 – neuronal excitability, 418 – structural basis for the restricted specificity, 413, 414 – plasmin, 411, 413, 414, 416 – urokinase-type plasminogen activator (uPA), 414, 415 – cell migration, 413, 416 – expression in central nervous system, 413 – potential natural substrates of uPA, 413, 414 – substrate specificity of uPA, 413, 414 tissue-type plasminogen activator (tPA) – activation of plasminogen by tPA, 413, 414 – axonal rearrangement, 416 – expression in central nervous system, 414 – long-term potentiation, 416–418 – migration of neurons, 416 – neurite outgrowth, 416, 427, 435 – synaptic plasticity, 416, 427

695

696

Index Protease, serine, 409 – activation domain, 411 – activation peptide, 418, 425 – biological functions of serine proteases, 432 – catalytic triad (His57, Asp102, Ser195), 412, 425, 432 – exon shuffling, 412, 414, 418 – mechanism of action, 416 – protease domain and additional structural domains, 411–414, 418, 425, 429–431 – signal peptide, 411, 418, 425 – specific subsites (SchechterBerger nomenclature), 412 – structural change during activation, 432 – substrate specificity, 412, 413, 418, 425, 433 – tertiary structure of the protease domain, 410 – active site, 411–413, 425, 432, 433 – b-barrel, 411, 425 – disulfide bridges, 412, 413 – oxyanion hole, 411, 432, 433 – substrate binding site, 412 Proteasomal activity, 656–659 Proteasomal subunits, 657, 659 20S proteasome, 377, 379, 382–385, 392, 393, 395, 396, 398 – a-type subunits, 374, 376, 378 – b-type subunits, 374–376 – chymotrypsinlike activity, 375 – N-terminal threonine nucleophile, 373, 375 – peptidylglutamyl peptide hydrolyzing activity, 375 – trypsinlike activity, 389 26S proteasome, 387–398 Proteasome and Parkinson’s disease, 383–390 – autosomal recessive juvenile parkinsonism, 373, 387 – CDCrel‐1, 387, 388 – Parkin, 386, 387 – Parkin-associated endothelin receptor-like receptor (Pael-R), 388 – phospholipase D, 385 – synaptogamin XI, 388 – a-synuclein, 389, 390 – synphilin‐1, 386, 389, 390 – tyrosine hydroxylase, 384, 389, 390 Proteasome inhibitor(s), 384, 391, 394, 398, 399, 657, 658 – epoxomicin, 385, 390

– lactacystin, 374, 385, 387, 389, 390, 392, 395, 396 – MG132, 375, 389 – Z-Ile-Glu(O-tBu)-Ala-leucinal (PSI), 373 Proteasome(s), 350, 351, 359, 361, 363, 558, 655, 657–660, 667, 669 – age-related changes, 665, 666 – assembly of, 666 – in centenarians, 666 – mouse mutations, 668 – ubiquitin proteasome system, 654 Protein – accumulation, 659 – cross links, 244, 254 – folding, 303 – synthesis, 170, 171, 173, 186–189 Proteinases activated receptor‐1 (PAR‐1), 568, 578, 579, 581 Protein phosphatase 1, 3, 18 Protein-protein interactions – caspase-recruiting domain (CARD), 518, 520, 529 – death domain (DD), 518, 525, 526, 529 – death effector domain (DED), 518, 519, 526 Proteoglycanase, 567 Proteoglycans, 285–287, 290, 294, 296–298, 568 – aggrecan, 566, 567, 569–571, 573 – brevican, 566, 571–573, 577 – chondroitin sulfate proteoglycans, 284, 297 – heparan sulfate proteoglycans, 284, 294, 298 – neurocan, 566 – versican, 566, 571, 573 Proteomics, 275, 276 Protofibrillary form, 660 Pruning, 656 PTPNS1, 42, 47, 50–52, 61–66 Punc, 42, 119, 120 Purkinje neurons, 668 – sensitivity to stress, 669 Puromycin, 626, 629, 635–637, 639, 642–644 PVR, 42, 66, 68, 69, 71–73 RAS suppressor, 275 Reactive oxygen species, 654, 658 Receptor tyrosine kinases, 656 Receptor, 269, 270 Regulated intramembrane proteolysis (RIP) – g secretase complex (gSC), 471, 475, 476, 483–493 – isolation, 485, 486, 495, 497

– specificity, 487 – inhibition, 192, 489, 490 – nicastrin (NCT) (aph‐2), 470, 471, 475, 483–485, 495 – aph‐1, 471, 483, 484, 491, 495, 496 – pen‐2, 471, 483–485, 491, 495 Regulated secretory pathway, 156, 158, 159 – in endocrine cells, 154, 155 – in neuroendocrine cells, 154 – sorting of neuropeptides and BDNF to, 155–157 – sorting receptors for, 155, 164 – sorting signals for, 155, 166 11S regulator, 376 19S regulatory complex, 377–379, 382, 394, 396 Renpenning syndrome, 265 Rett syndrome, 271, 276 Reversion-inducing cysteine-rich protein with Kazal motifs (RECK), 572 Rho GEF6, 268 Rho GTPases, 267, 268 Rho guanine nucleotide exchange factors, See Rho GEF Ribosome, 170–174, 180, 181, 187, 188 RNCAM, 42, 96 Robo, 42, 43, 97–101 ROS, 658 Rubinstein-Taybi syndrome, 268 19 S cap complex, 379 – m-particle, 377, 378 – AAA ATPase, 378 – PA700, 392 – regulatory particle triple A protein (Rpt), 378 – regulatory protein non-ATPase (Rpn), 378 – Rpn 1, 378 – Rpn 2, 378 – Rpn 10 (S5a), 378, 394 – Rpn 11, 378, 382 – Rpt 5 (S6´), 378, 385 Schizophrenia, 212, 215 SDK1, 42, 116–118 SDK2, 116–118 Secretases, 470–472, 475, 476, 478, 479, 492, 496, 497 a‐Secretases (K16L17 cleavage) – ADAM, 471, 475, 476, 478, 489–497 – TNF-a‐converting enzyme (TACE), 471 – characterization, 475, 476

Index – domains, 476, 477 – tissue inhibitors of metalloproteases (TIMP), 478 – therapeutics, 479 b-secretases, b-APP-converting enzymes (BACE), 479 – characterization, 480, 481 – domains, 480 – pharmacology, 481–483 – specificity, 481 Secretory granule proteins, 155 – degradation of, 157, 163, 164 – stability of, 163 Senile plaque, SP, 470 Serine protease inhibitors – a1-Antichymotrypsin (ACT), 592, 595 – antithrombin, 597, 598 – neuroserpin, 592, 597 – pigment epithelium-derived factor (PEDF), 598 – plasminogen activator inhibitor (PAI), 598 – protease nexin I [Glia-derived nexin (GDN)], 595, 596 – protease nexin II [Amyloid precursor protein (APP)], 594 Sheddase, 566, 577, 578 – ectodomain shedding, 569, 570, 581 SHPS‐1, 43, 61 Signal-regulatory proteins, 60 Signal transduction, 173, 179, 180, 182, 184, 185 Slit-Robo GTPase, 268 Smac/Diablo, 517, 534, 535 – endogenous inhibitor of IAPs, 520 – release from mitochondria in apoptosis, 520 Smad interacting, 271 Small nuclear ribonucleoprotein, 266 Smurf1, 656 SNP analysis, 275, 276 Sonic hedgehog, 269 Sorting receptor – carboxypeptidase E, 157–159 – for BDNF – for proinsulin, 155–157 – for proopiomelanocortin, 155 Sorting signal, 159 – acidic amino acids in, 154 – hydrophobic amino acids in, 155, 156 – in BDNF, 155 – in proinsulin, 155, 157, 158 – in proopiomelanocortin, 155

Spinal cord injury, 447, 448, 459 – calpain activation and neurodegeneration, 453 – calpain inhibitors for neuroprotection, 453 – devastating secondary injury process, 452, 453 – increases in intracellular free Ca2+ levels, 452, 453 – pathophysiology, 452, 453 – primary injury, 452 Spine apparatus, 172, 174 Spinocerebellar ataxia type 1, 659 Spinocerebellar ataxias, 668 – sporadic AD, SAD, 474 Stearyl-Nle17-VIP – SNV, 674, 675, 677 Stromelysins, 569 Substrate targeting for ubiquitination, 382 – cyclin, 381 – N-end rule, 381 – PEST sequence, 381 – phosphorylation, 381 Sulfatases, 285 – lysosomal, 287 – endo-sulfatases, 284, 287, 298 – SULF1, 294–296 – SULF2, 294–296 Sulfate channels, 286, 287 Sulfotopes, 284–287, 290, 298 – antithrombin binding site, 294 – growth-factor binding and sequestrtation, 295 – viral docking sites, 291 Sulfotransferase inhibitors, 293 Sulfotransferases, 292, 293 – cytosolic sulfotransferases, 288 – role in detoxification and steroid metabolism, 284, 287 – golgi-associated sulfotransferase, 284 SUMO, 654, 658 Synapse, 170–174, 178–189, 578 Synapse-associated polyribosome complex (SPRC), 170, 174 Synaptic plasticity, 659 Synaptodendrosomes, 173, 175, 178, 186 Synaptoneurosomes, 173, 175, 186 Syndecan, 566–578, 580 a-synuclein, 658 TAG‐1, 45, 47, 63, 75, 83, 91, 92, 103, 104, 107, 111, 112 Tandem mass spectrometry/mass spectrometry, See LC-MS/MS Targeting signal, 654

Tau, 682, 683 a-thalassemia, 266, 267 Therapy, 363, 365 Thrombin, 556–558 Thrombospondin, 566, 567, 570, 571 Thrombotic thrombocytopenia purpura, 570 Thymosin b, 226, 231, 238 – sequestering activity of, 230 Tick-borne encephalitis, 593 Tissue inhibitors of metalloproteases (TIMP), 552, 558, 567, 572 TMEM25, 43, 119, 120 Trafficking, 657, 658 Transcription, 356–359, 361–363, 365, 366 Transforming growth factor b, 410, 425 Transglutaminases, 244, 245 trans-interactions, 44, 47–49, 51, 56, 58, 65, 69–71, 75, 77, 79, 80, 82, 85, 88, 90–93, 99, 100, 104, 107, 110, 111, 115 Translation, 170–174, 176, 178, 180, 183–187, 189 Translation regulation, 3, 12, 16, 21 Transport, 170, 172–174, 178, 179, 182, 184, 190 Traumatic brain injury, 448 – Ca2+ channel blockers, 453 – diffuse brain injury, 454 – fluid-percussion brain injury, 454 – involvement of calpain and caspase‐3, 454 – loss of cytoskeletal proteins, 454 – pathophysiological pathways, 453 Tropomyosin, 232, 233 – caldesmon binding of, 234 – increase the stability of F-actin, 234 – inhibit the ADF/cofilin activity, 229, 234, 238 – inhibit the drebrin activity – inhibit the fascin activity, 234 – inhibt the gelsolin activity, 234 – inhibit the myosin activity, 235 – isoforms of, 234 Tumor necrosis factor, 569 Tumor necrosis factor converting enzyme (TACE), ADAM17, 569, 574, 578 Ubiquitin carboxy-terminal hydrolases, 655, 658 Ubiquitin elongation factor, E4 – CHIP, 373, 382, 388

697

698

Index – U-box protein, 382 – Hsc70, 373, 382 Ubiquitin hydrolase – deubiquitinating enzyme, 378, 382 – PGP 9.5, 382 – ubiquitin C-terminal hydrolase (UCH), 382 – UCHL‐1, 382 – ubiquitin specific processing protease (UBP), 382 Ubiquitin ligase, 654–659 Ubiquitin ligation pathways, 656 Ubiquitin pools, 668–670 – contribution to neuronal sensitivity, 667 Ubiquitin, 383, 385, 390–396, 654–659 – anaphase promoting complex, 381 – Angleman’s syndrome, 381 – HECT domain, 381 – ring finger, 381, 386 – SCF complex, 381 – ubiquitin activating enzyme, E1, 380, 382 – ubiquitin conjugating enzyme, E2, 381, 382, 394 – UbcH7, 386, 388 – UbcH8, 386, 388

– ubiquitin-protein ligase, E3, 373, 381, 387 – VBC complex, 381 Ubiquitin-activating enzyme, 654 Ubiquitination, 654–659 Ubiquitin-binding subunits, 655 Ubiquitin-conjugates, 656 Ubiquitin-conjugating enzymes, 654 Ubiquitin-independent proteolysis – cyclin-dependent kinase inhibitor p21cip1, 382 – microtubule associated protein tau, 383 – ornithine decarboxylase, 373, 379, 382 Ubiquitin-interacting motif – ubiquitin-binding protein, 391, 394 – ubiquitin conjugating enzyme E2‐25K, 394 – valosin-containing protein, 394 Ubiquitin-specific processing enzymes, 655 UBR1, 656 UCH, 655, 658 UCH-L1, 658, 669 – mutation in mouse, 668

Un-translated region (UTR), 170, 181, 182, 184, 189 Unverricht-Lundborg disease, 603 UPS, 654–656, 658–660 USP14, 670 – mutation in mouse, 668 Vascular dementia, 605 Vasoactive intestinal peptide (VIP), 674–677 Voltage-gated Ca2+ channels – Ca2+ translocation across the plasma membrane, 450 – L-type channels, 450 – membrane depolarizations, 450 – Na+/K+-ATPase, 449, 450 – transmembrane ionic gradients, 450 – T-type channels, 450 Voltage-gated ion channels, 449 – alteration in membrane potential, 450 – conformational changes, 450 – voltage-dependent activation, 450 von Willebrand factor Proteinase, 567 WAVE, actin polymerization promoting activity of, 231

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Handbook of Neurochemistry and Molecular Neurobiology: Schizophrenia (Springer Reference)

Handbook of Neurochemistry and Molecular Neurobiology - Neurotransmitter Systems

Handbook of Neurochemistry and Molecular Neurobiology, 3rd Edition Neuroimmunology

Handbook of Neurochemistry and Molecular Neurobiology: Neural Signaling Mechanisms (Springer Reference)

Handbook of Neurochemistry and Molecular Neurobiology 3rd Edition: Practical Neurochemistry Methods

Handbook of Neurochemistry and Molecular Neurobiology 3rd Edition: Sensory Neurochemistry (Springer Reference)

Handbook of Neurochemistry and Molecular Neurobiology (Amino Acids and Peptides in the Nervous System)

Cellular and Molecular Neurobiology

Protein stucture and function

Protein structure and function

Protein structure and function

Cellular and Molecular Neurobiology,

Neural Networks and Brain Function

Elements of Molecular Neurobiology

Elements of Molecular Neurobiology

Elements of Molecular Neurobiology

Elements of Molecular Neurobiology

Roots of Social Sensibility and Neural Function

Cyclic GMP: Synthesis, Metabolism, and Function

Protein Metabolism and Homeostasis in Aging

A Handbook of Bioanalysis and Drug Metabolism

Handbook of drug metabolism

A Handbook of Bioanalysis and Drug Metabolism

Mitochondrial Function: Mitochondrial Protein Kinases, Protein Phosphatases and Mitochondrial Diseases

Protein Stucture and Function (Primers in Biology)

Handbook of Neurochemistry and Molecular Neurobiology. Neural Protein Metabolism and Function