Amyloid Precursor Protein A Practical Approach

Author: Weiming Xia | Huaxi Xu

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Amyloid Precursor Protein A Practical Approach

© 2005 by CRC Press LLC

Amyloid Precursor Protein A Practical Approach EDITED BY

Weiming Xia and Huaxi Xu

CRC PR E S S Boca Raton London New York Washington, D.C.

© 2005 by CRC Press LLC

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Library of Congress Cataloging-in-Publication Data Xia, Weiming. Amyloid precursor protein : a practical approach / by Weiming Xia, Huaxi Xu. p. cm. Includes bibliographical references and index. ISBN 0-8493-2245-6 (alk. paper) 1. Amyloid beta-protein precursors — Laboratory manuals. I. Xu, Huaxi. II. Title. QP552.A45X53 2004 616.8′047—dc22

2004058142

This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage or retrieval system, without prior permission in writing from the publisher. All rights reserved. Authorization to photocopy items for internal or personal use, or the personal or internal use of specific clients, may be granted by CRC Press, provided that $1.50 per page photocopied is paid directly to Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923 USA. The fee code for users of the Transactional Reporting Service is ISBN 0-8493-2245-6/05/$0.00+$1.50. The fee is subject to change without notice. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. The consent of CRC Press does not extend to copying for general distribution, for promotion, for creating new works, or for resale. Specific permission must be obtained in writing from CRC Press for such copying. Direct all inquiries to CRC Press, 2000 N.W. Corporate Blvd., Boca Raton, Florida 33431. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation, without intent to infringe.

Visit the CRC Press Web site at www.crcpress.com © 2005 by CRC Press No claim to original U.S. Government works International Standard Book Number 0-8493-2245-6 Library of Congress Card Number 2004058142 Printed in the United States of America 1 2 3 4 5 6 7 8 9 0 Printed on acid-free paper

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Preface Amyloid precursor protein (APP) is an extensively studied single transmembrane protein. Our vast knowledge of this protein is derived from thousands of reports published during the past decade. In fact, many scientists from different disciplines have used their well-established experimental procedures to examine the characteristics of this molecule. Hence, their published reports display nearly all aspects of biological techniques used in genetics, molecular biology, cell biology, and biochemistry. As a result, APP may be viewed as a unique model protein to illustrate a wide array of basic and advanced biological techniques used in many laboratories. The major aim of this book is to demonstrate the critical techniques utilized in the experiments selected from many significant findings that contribute to our understanding of APP biology. Each technique will be presented in the format of a standard protocol, providing step-by-step instructions for bench scientists carrying out similar studies on APP and other proteins. Theoretical background and discussions will be provided in the introduction section, and a brief description of the goal will also be presented. These protocols will form the core of our approach in elucidating the function of a protein. The antibodies used in each experiment will be described and are cited on the list of antibodies to APP and Aβ proteins at the front of this book It is our intention to contrast basic and advanced methods and demonstrate how development of biological techniques significantly affects the way we examine our molecular targets. An important feature of this book is the presentation of modifications applied to standard procedures used in examining a membrane protein. These modifications will likely help readers consider similar alterations in their own experimental procedures. The fact that most experiments we perform every day do not lead to conclusive answers strongly indicates that actively modifying our approach is essential to perform biological experiments and achieve definitive results. The descriptions of the modifications will help justify similar alterations in readers’ own experimental approaches. Another goal of this book is to include commonly used experimental procedures and clearly present them in the format of a protocol to serve as a laboratory manual for bench scientists working on different aspects of the biological functions of APP and other membrane proteins. Most experimental procedures can be carried out in a regularly equipped laboratory, but a few protocols require sophisticated core facilities. In addition, multistep protocols will be broken down into several independent protocols, allowing an investigator to create parallel experiments to accelerate the achievement of results. This book will also describe a set of previously published milestone studies on APP. Results summarized here will not only provide a complete picture of our current understanding of APP, but also confer future direction for continued investigation of this protein in normal cellular function and in disease.

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Due to rapid expansion of our knowledge of APP biology, this book will cover the most up-to-date research activities. More importantly, we emphasize practical techniques used to address key questions related to APP and similar membrane proteins. Hence, this book will offer a framework for studying other membrane proteins and provide detailed, step-wise procedures to achieve specific aims. It will be suitable for students who are learning basic experimental approaches to address biological questions and also for bench scientists who seek immediate assistance and practical approaches for studying proteins.

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Contributors Karen Hsiao Ashe Department of Neurology University of Minnesota Minneapolis, Minnesota

Erica A. Fradinger Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Jorge Busciglio Department of Neurobiology and Behavior University of California Irvine, California

Denise Galatis Department of Pathology The University of Melbourne Melbourne, Victoria, Australia

Dongming Cai Fisher Center for Research on Alzheimer’s Disease The Rockefeller University New York, New York

Arun K. Ghosh Department of Chemistry University of Illinois at Chicago Chicago, Illinois

William A. Campbell Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts Roberto Cappai Department of Pathology The University of Melbourne Melbourne, Victoria, Australia Atul Deshpande Department of Neurobiology and Behavior University of California Irvine, California Susanne C. Feil Biota Structural Biology Laboratory St. Vincent’s Institute of Medical Research Fitzroy, Victoria, Australia

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Gunnar Gouras Department of Neurology and Neuroscience Weill Medical College of Cornell University New York, New York Heike S. Grimm Center for Molecular Biology University of Heidelberg Heidelberg, Germany Marcus O.W. Grimm Center for Molecular Biology University of Heidelberg Heidelberg, Germany Tobias Hartmann Center for Molecular Biology University of Heidelberg Heidelberg, Germany

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Pablo Helguera Department of Neurobiology and Behavior University of California Irvine, California

Samir Kumar Maji Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Stanley Jones Premkumar Iyadurai Department of Neurology University of Minnesota Minneapolis, Minnesota

William J. McKinstry Biota Structural Biology Laboratory St. Vincent’s Institute of Medical Research Fitzroy, Victoria, Australia

Gerald Koelsch Protein Studies Program Oklahoma Medical Research Foundation Oklahoma City, Oklahoma

Chica Mori Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Edward H. Koo Department of Neurosciences University of California, San Diego La Jolla, California

William J. Netzer Fisher Center for Research on Alzheimer’s Disease The Rockefeller University New York, New York

Markus P. Kummer Department of Neurosciences University of California, San Diego La Jolla, California Noel D. Lazo Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Andreas J. Paetzold Center for Molecular Biology University of Heidelberg Heidelberg, Germany Michael W. Parker Biota Structural Biology Laboratory St. Vincent’s Institute of Medical Research Fitzroy, Victoria, Australia

Cynthia A. Lemere Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Alejandra Pelsman Department of Neurobiology and Behavior University of California Irvine, California

Feng Li Center for Neuroscience and Aging The Burnham Institute La Jolla, California

Thomas Ruppert Center for Molecular Biology University of Heidelberg Heidelberg, Germany

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Dongwoo Shin Department of Chemistry University of Illinois at Chicago Chicago, Illinois

Vajira Weerasena Protein Studies Program Oklahoma Medical Research Foundation Oklahoma City, Oklahoma

Xiaoyan Sun Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Michael S. Wolfe Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Reisuke H. Takahashi Department of Neurology and Neuroscience Weill Medical College of Cornell University New York, New York

Weiming Xia Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Akihiko Takashima Brain Science Institute Institute of Physical and Chemical Research Saitama, Japan

Huaxi Xu Center for Neuroscience and Aging The Burnham Institute La Jolla, California

Jordan Tang Protein Studies Program Oklahoma Medical Research Foundation Oklahoma City, Oklahoma

Tsuneo Yamazaki Department of Neurology Graduate School of Medicine Gunma University Gunma, Japan

David B. Teplow Center for Neurologic Diseases Brigham and Women’s Hospital Harvard Medical School Boston, Massachusetts

Hui Zheng Department of Molecular and Human Genetics Baylor College of Medicine Houston, Texas

Eva G. Zinser Center for Molecular Biology University of Heidelberg Heidelberg, Germany

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Table of Contents List of Antibodies to APP and Aβ Proteins Chapter 1

Biochemical Characterization of Amyloid Precursor Protein

Weiming Xia Chapter 2

Assays for Analysis of APP Secretion and Recycling

Markus P. Kummer, Tsuneo Yamazaki, and Edward H. Koo Chapter 3

Strategies for Crystallizing the N-Terminal Growth Factor Domain of Amyloid Precursor Protein

William J. McKinstry, Susanne C. Feil, Denise Galatis, Roberto Cappai, and Michael W. Parker Chapter 4

Analysis of Amyloid Precursor Protein Processing Protease β-Secretase: Tools for Memapsin 2 (β-Secretase) Inhibition Studies

Gerald Koelsch, Vajira Weerasena, Dongwoo Shin, Arun K. Ghosh, and Jordan Tang Chapter 5

Assays for Amyloid Precursor Protein γ-Secretase Activity

William A. Campbell, Michael S. Wolfe, and Weiming Xia Chapter 6

Cell-Free Reconstitution of β-Amyloid Production and Trafficking

Dongming Cai, William J. Netzer, Feng Li, and Huaxi Xu Chapter 7

Studying Amyloid β-Protein Assembly

Erica A. Fradinger, Samir Kumar Maji, Noel D. Lazo, and David B. Teplow Chapter 8

Intracellular Accumulation of Amyloid β and Mitochondrial Dysfunction in Down’s Syndrome

Jorge Busciglio, Alejandra Pelsman, Pablo Helguera, and Atul Deshpande

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Chapter 9

Linking Alzheimer’s Disease, β-Amyloid, and Lipids: A Technical Approach

Marcus O.W. Grimm, Andreas J. Paetzold, Heike S. Grimm, Eva G. Zinser, Thomas Ruppert, and Tobias Hartmann Chapter 10 Regulation of Amyloid Precursor Protein Processing by Lithium Xiaoyan Sun and Akihiko Takashima Chapter 11 Immunocytochemical Analysis of Amyloid Precursor Protein and Its Derivatives Gunnar Gouras and Reisuke H. Takahashi Chapter 12 Pathological Detection of Aβ and APP in Brain Chica Mori and Cynthia A. Lemere Chapter 13 Creating APP Transgenic Lines in Mice Stanley Jones Premkumar Iyadurai and Karen Hsiao Ashe Chapter 14 Generation of Amyloid Precursor Protein Knockout Mice Hui Zheng

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List of Antibodies to APP and Aβ Proteins Name

Source

Immunogen (Specific)

22C11

Chemicon Cat# MAB348 N terminal, purified recombinant APP A4 fusion protein

Alz-90 (1.D5)

Chemicon Cat# MAB349 Synthetic aa 511-608 of APP pre A4-695

369

S. Gandy C terminal APP [email protected] Elan Pharmaceuticals Specific to carboxyl terminus of β-secretase cleavage site of APP D. Selkoe Residues 595–611 of [email protected]. APP695 (α-sAPP) harvard.edu D. Selkoe Against 20 C terminal [email protected]. residues of APP harvard.edu D. Selkoe Against 20 C terminal [email protected]. residues of APP harvard.edu David Miller Residues 672–695 of [email protected] APP695 Elan Pharmaceuticals Residues 444–591 of APP (β−APPs and APP) Elan Pharmaceuticals Residues 676–695 of APP695 E.H. Koo Residues 380–665 of APP [email protected] EH. Koo Recognize [email protected] nonoverlapping epitopes in extracellular region of APP

APP 192

1736

C7

C8

R57 8E5 13G8 1G7 5A3

© 2005 by CRC Press LLC

Host Description (Formulation) Methods in Chapter Monoclonal mouse (lyophilized, azide) Monoclonal mouse (purified, lyophilized, azide) Polyclonal

IH, WB

6

WB

8

IH, WB

6

Polyclonal

IH, WB

1

Polyclonal

WB

1

Polyclonal

IH, IP

1, 5, 12

Polyconal

IP, WB

8

Polyclonal

WB

5

Monoclonal

WB

12

Monoclonal

IP, WB

1, 5

Monoclonal

IP, IF, WB IP, IF, WB

2

Monoclonal

2

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4G8

Signet Cat# 9200, 9220

6E10

Signet Cat# 9300, 9320

1280

D. Selkoe [email protected]. harvard.edu Genetics Company Cat# AB-10

W0-2

G2-10

Genetics Company Cat# AB-10

G2-11

Genetics Company Cat# AB-11

MBC40

β-amyloid 1-42

H. Yamaguchi [email protected] H. Yamaguchi [email protected] Chemicon Cat# AB5078P

21F12

Elan Pharmaceuticals

R1282

D. Selkoe [email protected]. harvard.edu

MBC42

Amino acid residues Monoclonal, 17–24 of β-amyloid mouse peptide (reactive to aa (crude, 17–24 Aβ and to APP) ascites) Amino acid residues 1–17 Monoclonal, of β-amyloid peptide mouse (reactive to aa 1–17 Aβ (crude, and to APP) ascites) Raised to Aβ 1–40 Polyclonal (reactive to Aβ and P3) aa 1–10 of N terminal of human Aβ (high affinity to amyloid peptides Aβ1-38, Aβ1-39, Aβ1-40, Aβ1-43, and Aβ1-44) Amino acid residues 31–40 of human Aβ peptide at C terminal (Aβ-peptide, aa 31–40; not Aβ1-38, Aβ1-39, Aβ1-42, Aβ1-43, or Aβ1-44) Amino acid residues 33–42 of human Aβ 42 peptide at C terminal (Aβ-peptide, aa 33–42; not Aβ1-38, Aβ1-39, Aβ1-40, Aβ1-43, or Aβ1-44) Amino acid residues 1–40 of β-amyloid peptide (reactive to Aβ 40) Amino acid residues 1–42 of β-amyloid peptide (reactive to Aβ 42) β-amyloid 1-42 (recognizes β-amyloid 1-42) β-amyloid 33-42

Aβ 1−40

ELISA, IH, IP, WB

6, 12

ELISA, IH, IP, WB

6, 10, 12

IP

1

Monoclonal mouse

ELISA, WB

9

Monoclonal mouse

ELISA, WB

9

Monoclonal mouse

ELISA, WB

9

Monoclonal

IH, WB

10, 11, 12

Monoclonal

IH, WB

10, 11, 12

Polyclonal rabbit

WB, IH, IF, IP, ELISA IH, ELISA, WB IH, IP

11

Monoclonal mouse Polyclonal rabbit

12

12

IH = Immunohistochemistry; WB = Western blot; IP = Immunoprecipitation; IF = Immunofluorescence; ELISA = Enzyme-linked immunosorbent assay.

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1

Biochemical Characterization of Amyloid Precursor Protein Weiming Xia

CONTENTS 1.1 1.2 1.3 1.4 1.5

Introduction Main Scheme of Approaches Results Discussion Protocols 1.5.1 Overexpression of APP in Mammalian Cells by Transient Transfection 1.5.2 Selection of Stable Cell Line Overexpressing APP 1.5.3 Determination of Protein Concentration by BCA 1.5.4 Identification of APP and Its Derivatives by Western Blot 1.5.5 Radiolabeling of Cells with [35S]-Met 1.5.6 Identification of Full-Length APP and Its Derivatives by Immunoprecipitation 1.5.7 Determination of Half-Life of APP 1.5.8 Co-Immunoprecipitation of APP-Interacting Protein 1.5.8.1 Preparation 1.5.8.2 Pre-Absorption of Protein A-Sepharose 1.5.8.3 Pre-Clearing of Cell Lysates 1.5.8.4 Set Up Co-IP 1.5.8.5 Wash Co-IP 1.5.9 Conjugation of Antibody to Protein A-Sepharose Acknowledgments References

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1.1 INTRODUCTION Amyloid precursor protein (APP) is a single transmembrane protein that undergoes sequential proteolysis to generate multiple peptides, including the amyloid β-peptide (Aβ) — the major component of the senile plaques that are diagnostic hallmarks of Alzheimer’s disease (AD).1,2 AD accounts for more than 50% of cases of dementia in the elderly and has a prevalence estimated at 15 to 20 million patients worldwide. It is associated with progressive memory loss that leads to profound dementia and eventually death, although a patient can have the disease for as long as 10 to 15 years before death. The pathology of AD is characterized by extracellular neuritic plaques consisting of Aβ and intracellular neurofibrillary tangles.3 A central role for Aβ in the pathogenesis of AD was first discovered by finding APP mutations in a subset of familial AD (FAD) cases that occurred as inherited autosomal dominant disease.4,5 The APP gene is located on chromosome 21, and mutations found in APP occur either within the Aβ peptide sequence (A692G6 and E693Q,7,8 APP770 numbering) or immediately flanking the Aβ peptide sequence including KM670/671NL (Swedish mutation),9 I716V,10 and V717I,G,F mutations.5,11,12 Mutations in APP are rare but they have been very informative. Patients carrying trisomy 21 (Down’s syndrome) develop the histopathology of AD in early midlife, presumably because they have three copies of the APP gene and a documented increase in APP transcription13 that leads to augmented Aβ deposition. The double mutation in APP at the Aβ N terminus (Swedish mutation) leads to a marked increase in Aβ production,9,14–16 as confirmed in primary skin fibroblasts and the plasma of presymptomatic and symptomatic carriers.17 The mutations at APP716 or 7175,11,12 lead to hypersecretion of the longer and more amyloidogenic Aβ42 peptide.10,18 When human APP containing the Val → Phe mutation19 or the Swedish mutation20 is overexpressed in transgenic mice, a time-delayed accumulation of both diffuse and neuritic Aβ plaques develops. These various studies provide strong circumstantial evidence for the early mechanistic role of abnormal APP metabolism and Aβ deposition in AD neuropathology. In this chapter, we will discuss several basic biochemical approaches routinely used to study full-length APP and shorter peptides derived from proteolytic cleavages of APP holoprotein (Figure 1.1); the same approaches can be used to characterize any newly identified proteins.

1.2 MAIN SCHEME OF APPROACHES To characterize a newly cloned gene product, transient transfection is a quick and simple way to enrich the protein of interest for biochemical analysis. Here we present a standard protocol to transiently transfect a mammalian expression vector containing APP cDNA into Chinese hamster ovary (CHO) cells (Protocol 1.5.1). During the 24 to 48 hr post-transfection, cells can either be harvested for immediate analysis or used for screening of clones to make a cell line stably expressing APP. In the latter case, the selection drug (based on the selection marker in the expression vector used for transient transfection) will be used for screening (Protocol 1.5.2).

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FIGURE 1.1 Proteolytic cleavage of amyloid precursor protein by α-, β- and γ-secretases. APP is proteolytically processed by two alternative pathways. First, α-secretase cleaves slightly N terminal to the beginning of the APP transmembrane domain (at residues 16 and 17 of the Aβ region) and generates a major secreted derivative (α-APPs) and a ~10-kDa C-terminal fragment (C83). C83 can be cleaved by a protease activity called γ-secretase to yield p3. Alternative cleavage of APP by the β-secretase (called BACE or memapsin 2) generates a soluble N-terminal fragment (β-APPs) and a 12-kDa C-terminal fragment of APP (C99) that can be further cleaved by γ-secretase to yield two major species of Aβ ending at residue 40 (Aβ40) or 42 (Aβ42).

A large number of antibodies against different regions of APP are available from many laboratories and commercial sources. Using these antibodies, cell lysates with equivalent amounts of protein concentrations (determined by Bicinchoninic Acid Kit [BCA]; Protocol 1.5.3) can be separated by SDS-PAGE followed by Western blotting analysis (Protocol 1.5.4). An alternative approach would be to metabolically label cells with [35S]-methionine (Met) (Protocol 1.5.5), and lyse the radiolabeled cells for immunoprecipitation with specific antibodies (Protocol 1.5.6). Radiolabeling cells followed by immunoprecipitation usually enhances the signal-to-noise ratios of overexpressed proteins. Proteins identified by Western blot and immunoprecipitation usually represent mature species at steady state levels. To examine the process of protein maturation, cells can be pulse-labeled with [35S]-Met followed by chasing in nonradioactive media for various periods of time (Protocol 1.5.7). Cell lysates will then be immunoprecipitated with specific antibodies. A simple co-immunoprecipitation method using an antibody against a candidate protein (e.g., presenilin) can be used to determine whether this protein interacts with APP (Protocol 1.5.8). By using free antibody or protein A sepharose-conjugated

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antibody (Protocol 1.5.9), co-immunoprecipitation is widely used to confirm a candidate protein that specifically interacts with APP.

1.3 RESULTS APP occurs in three alternatively spliced forms of 695, 751, and 770 residues, and these proteins undergo N- and N + O glycosylation as well as phosphorylation. In CHO cells stably expressing APP, both N- and N + O-glycosylated APP holoproteins can be detected by immunoprecipitation of radiolabeled cells with antibody C7, which was raised against the last 21 amino acids of the APP C terminus (APP675–695, APP695 numbering; Figure 1.2a). APP is proteolytically processed by at least two broad alternative pathways (Figure 1.1). First, cleavage of APP by β-secretase (called BACE or memapsin 2)21–24 generates a soluble N-terminal fragment (β-APPs, Figure 1.2b), and a C-terminal stub of APP (C99, Figure 1.2d),25 which can be further cleaved by a protease activity called γ-secretase to yield two major species of Aβ ending at residue 40 (Aβ40) or 42 (Aβ42). See Figure 1.2e.26,27 Since soluble β-APPs and Aβ generated in radiolabeled cells are secreted into media, they can be detected by immunoprecipitation of media with

a.

b.

c.

100

100 β-APPs

N+O-APP

α-APPs

N-APP 100

d. 14

e. 6 C99 C83

Aβ 3

p3

FIGURE 1.2 Detection of APP and its derivatives by immunoprecipitation. (a) Both N- and N+O-glycosylated APP holoproteins were immunoprecipitated from radiolabeled CHO cells stably expressing APP with antibody C7, which was raised against the last 21 amino acids of the APP C terminus. (b) Soluble β-APPs from radiolabeled cells were secreted into media and could be detected by immunoprecipitation of media with specific antibody 192 which specifically recognizes the C termini of β-APPs. (c) Soluble α-APPs can be detected by immunoprecipitation of growth media with antibody 1736 which recognizes the C termini of α-APPs. (d) APP-expressing cells were lysed and immunoprecipitated with antibody C7, followed by Western blotting with another C terminus antibody, 13G8, to detect C99 and C83. (e) Both Aβ and p3 were immunoprecipitated from growth media of radiolabeled APPexpressing CHO cells using antibody 1280, which was raised against the whole region of Aβ.

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specific antibodies. Antibody 192 can specifically recognize the C termini of β-APPs (Figure 1.2b). Antibody 1280 was raised against the whole region of Aβ, and can thus recognize both Aβ and p3 (Figure 1.2e). Alternative cleavage slightly N terminal to the beginning of the APP transmembrane domain (at residues 16–17 of the Aβ region) by a α-secretase protease28,29 generates the major secreted derivatives, α-APPs (Figure 1.2c), precluding Aβ formation (nonamyloidogenic).30,31 The ~10-kDa C-terminal stub (C83, Figure 1.2d) can be cleaved by γ-secretase to yield p340 and p342 (Figure 1.2e),32 which can be recognized by antibody 1280. Soluble α-APPs can be detected by immunoprecipitation of growth media with antibody 1736, which recognizes the C termini of α-APPs (Figure 1.2c). Another approach to detecting APP and its derivatives is to perform immunoprecipitation followed by Western blotting; this approach does not require radiolabeling of cells. For example, APP-expressing cells can be lysed and immunoprecipitated with antibody C7. The immunoprecipitates, full length APP, C83, and C99, all carry the antigen for C7, namely, the C terminus of APP. After immunoprecipitates are separated by SDS-PAGE, Western blotting with another C-terminal antibody, 13G8, can be used to detect these peptides. This is a convenient approach to detect full length APP as well as C99 and C83 (Figure 1.2d). The temporal events of APP glycosylation and proteolytic processing occur within a very short period. Within a half hour of protein translation, the majority of APP is glycosylated, as determined by pulse-chase labeling of APP expressing cells with [35S]-Met (Figure 1.3). After APP-expressing cells were incubated with Metfree media for 45 min at 37˚C, cells were pulse-labeled with [35S]-Met for 5 min, and then the medium was changed to regular Dulbecco’s modified Eagle’s medium and chased for 0.25 to 5 hr. Cells were lysed and immunoprecipitated with antibody C7, followed by SDS-PAGE. The half life of APP was very short (~30 min) and most of the holoprotein was either degraded or proteolytically cleaved by β- or α-secretase to generate C99/C83 within 1.5 hr (Figure 1.3). Like the APP holoprotein, almost all of the C83/C99 was either degraded or cleaved by γ-secretase, and no C83/C99 was detectable after 5 hr (Figure 1.3). To determine the spatial distribution of immature and mature APP, membrane vesicles enriched in different subcellular compartments were separated by fractionation on discontinuous Iodixanol sucrose gradients (see Chapter 5). A total of 12 fractions were collected, and each fraction was analyzed by Western blotting. For endoplasmic reticulum (ER)-rich fractions, an antibody against the ER marker protein, calnexin, was used (Figure 1.4a). The densest fractions (1 through 4) had the strongest immunoreactivities for calnexin, indicating that these fractions contained ER vesicles. For Golgi/trans-Golgi network (TGN)-enriched fractions, β-1,4-galactosyltransferase activity in each fraction was measured, i.e., the addition of [3H]-galactose onto the oligosaccharides of an acceptor protein, ovomucoid, was measured.33 Since β-1,4-galactosyltransferase is a marker for Golgi/TGN-type vesicles, fractions 4 through 8 were enriched in Golgi/TGN vesicles (Figure 1.4b). Alternatively, an

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0

15

30

90 120

300 min

210 N+O-APP N-APP 110 71

28 18 14

C99 C83

6

FIGURE 1.3 Maturation of APP and turnover of APP and C terminal fragments. CHO cells stably expressing APP were pulse-labeled for 5 min and chased for 0 to 5 hr. Cell lysates were immunoprecipitated with C7. The half life of APP was ~30 min, and a portion of APP holoprotein was cleaved by β- or α-secretase to generate C99/C83 within 1.5 hr. Most C83 and C99 were either degraded or cleaved by γ-secretase, and no C83/C99 was detectable after 5 hr.

antibody against another Golgi/TGN marker protein, syntaxin 6, can be used to characterize these subcellular fractions. When these subcellular fractions were probed for APP distribution with APP monoclonal antibodies 5A3/1G7, which can detect both N- and N + O-glycosylated APP proteins (Figure 1.4c), fractions 1 through 3 only had N-glycosylated APP proteins. The lack of further glycosylation of these immature APP proteins suggests that N-glycosylated APP resides primarily in the ER (Figure 1.4c).34–36 Both N- and N + O-glycosylated APP proteins were observed from fraction 4 to fraction 8, indicating that post-translational modification is completed during the passage of APP into the Golgi/TGN compartment.34 Post-translationally modified APP continues to transport through the central vacuolar pathway and finally reaches the cell surface. A large portion of APP undergoes endocytosis and is re-internalized to endosomes (see Chapter 2). To examine whether another AD-linked gene product, presenilin 1 (PS1), interacts with APP, co-immunoprecipitation of APP and PS1 was carried out (Protocol 1.5.8). When cells were lysed and lysates were immunoprecipitated with PS1 antibodies X81, R22, or 4627 (against N-terminal, middle, and C-terminal regions of PS1, respectively) followed by Western blotting with the APP C-terminal antibody 13G8, full-length APP was clearly detected (Figure 1.5). The specificity of this interaction was demonstrated by the observation that only the N-glycosylated form of full-length APP co-precipitated with PS1 (Figure 1.5).

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a.

200

97

calnexin

b.

c.

Galactosyltransferase Activity (cpm)

68

800 600 400 200 0

200 N+O-APP N-APP 97 68 1

2

3

4

5

6

7

8

9

10

11 12

FIGURE 1.4 Characterization of immature and mature APP in subcellular fractions. (a) Distribution of the ER marker protein (calnexin) in discontinuous Iodixanol gradient fractions was detected by Western blotting with anti-calnexin antibody (densest fraction, lane 1; lightest fraction, lane 12). (b) Distribution of the Golgi/TGN marker β-1,4-galactosyl transferase activity was mainly in fractions 4 through 8. (c) The same fractions were immunoblotted with APP monoclonal antibodies 5A3/1G7. Fractions 1 through 3 were rich in ER vesicles and contained solely N-glycosylated APP; fractions 4 through 8 were rich in Golgi/TGN vesicles which contained both N- and N + O-glycosylated APP. Fraction 4 represented a transition fraction in the discontinuous gradient and contained both ER and Golgi/TGN proteins. (From Xia, W. et al. Proc. Natl. Acad. Sci. USA, 97, 9299–9304, 2000; Xia, W. et al. Biochemistry 37, 16465–16471, 1998. With permission.)

1.4 DISCUSSION This chapter has presented several basic approaches to characterize the biochemical properties of APP. These experimental procedures are routinely utilized in many laboratories, and the protocols listed in this chapter usually serve as starting methods to study transmembrane proteins. Various modifications can be introduced to meet special needs for individual proteins. While more than a dozen transfection reagents are available to introduce expression vectors into mammalian cells, the toxicity of reagents usually counteracts the transfection efficiency. Therefore, it is necessary to titrate the amount of DNA/transfection reagent to obtain the optimal transfection efficiency with minimum toxicity. A simple test is to co-transfect the gene of interest with another vector expressing green fluorescent protein (GFP), and the expression levels of GFP can be monitored under a fluorescent microscope 24 to 48 hr post-transfection. If cellular toxicity is

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PS1 antibody X81

4627

APP antibody R22

Preimm

C7

N+O-APP N-APP 110

1

2

3

4

5

FIGURE 1.5 Co-immunoprecipitation of APP and PS1. Lysates from APP- and PS1-expressing cells were co-immunoprecipitated with either PS1 antibodies (X81, 4627, or R22) or preimmune serum (preimm), followed by Western blotting with APP antibody 13G8. The APP species that co-immunoprecipitated with PS1 co-migrated with the N-glycosylated form of APP detected on straight Western blots of the lysates. The lower band in lanes 2 through 4 is nonspecific. (From Xia, W. et al. Proc. Natl. Acad. Sci. USA, 97, 9299–9304, 2000. With permission.)

obvious even when a low amount of DNA/transfection reagent is used, then overexpressing the target gene may cause enough cellular damage that raising a stable cell line is not feasible. Detection of proteins by immunoprecipitation and/or Western blot largely depends on the specificity of an antibody. Nevertheless, the choice of detergent in the lysis buffer is important to successfully lyse a membrane protein. This is especially critical when co-immunoprecipitation is performed to search for any interacting proteins that form a complex with the transmembrane protein. Because many transmembrane proteins tend to interact nonspecifically and form aggregates under non-physiological conditions, i.e., overexpression of two transmembrane proteins in transiently transfected cells, stringent conditions should be tested to differentiate a specific protein–protein interaction versus nonspecific hydrophobic aggregation. In some cases, steric hindrance may interfere with an antibody binding to a specific region during co-immunoprecipitation, Therefore, using multiple antibodies against different regions of the protein for co-immunoprecipitation is necessary to confirm a specific interaction. Reverse co-immunoprecipitation should be carried out to prove a direct interaction between two proteins. In addition, examining the occurrence and localization of the complex will also help explain the physiological significance of the interaction between two proteins. In conclusion, the standard protocols listed in this chapter provide an outline of experiments that can be immediately performed to study a new protein. Understanding the basic biochemical properties of a protein is not only the basis for future in vitro and in vivo studies, but also represents the first step to explore its biological function.

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1.5 PROTOCOLS 1.5.1 OVEREXPRESSION OF APP IN MAMMALIAN CELLS BY TRANSIENT TRANSFECTION 1. Split CHO cells in a six-well plate the day before transfection so that they are 90 to 95% confluent the following day. Plate cells in 1.5 ml of their normal growth medium. 2. For each well of cells, dilute 4 µg of DNA in 250 µl medium without serum (e.g., Opti-MEM I). Additionally, dilute 12 µl Lipofectamine 2000 (Invitrogen, #11668-019) in 250 µl Opti-MEM I for each well of cells and incubate for 5 min at room temperature. 3. Combine diluted Lipofectamine and the DNA within 30 min. Incubate at room temperature for 20 min to allow DNA–Lipofectamine complexes to form. 4. Add 500 µl of the DNA–Lipofectamine mixture to each well of cells and mix gently. 5. Incubate cells at 37˚C in a CO2 incubator for 24 to 48 hr. Growth medium may be replaced after 4 to 6 hr.

1.5.2 SELECTION

OF

STABLE CELL LINE OVEREXPRESSING APP

1. At 48 hr post-transfection, detach the cells by brief trypsin treatment, count the cells, and make serial dilutions to obtain a final concentration of 1 cell/100 µl of media containing a selection drug (e.g., G418). Transfer 100 µl of media to each well of a 96-well plate, resulting in a final cell count of approximately one cell per well. A total of six to eight plates should be prepared for selection of multiple clones of the stable cell line. 2. Formation of a single colony of cells in individual wells is monitored under a microscope after a growth period of 2 weeks. Only wells containing single colonies of cells will be selected. Transfer cells to a 24-well plate for growth. 3. Prepare duplicate wells of cells from the same clone and lyse one well of cells for measuring expression levels of APP by Western blot. 4. Any candidate clones with satisfactory expression levels of APP will be grown in large quantities for long-term storage.

1.5.3 DETERMINATION

OF

PROTEIN CONCENTRATION

BY

BCA

1. Prepare BSA standards in lysis buffer [50 mM Tris, pH 7.6, 150 mM NaCl, 2 mM EDTA, 1% NP-40 and a protease inhibitor cocktail (5 µg/ml leupeptin, 5 µg/ml aprotinin, 2 µg/ml pepstatin A, and 0.25 mM phenylmethylsulfonyl fluoride)] so that there is a serial dilution of BSA from 1 mg/ml down to 15.5 µg/ml. 2. In a 96-well plate, add 25 µl of sample or standard to each well.

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3. Mix 50 parts of solution A to 1 part of solution B from the BCA Protein Assay Reagent (Pierce, #23225). Add 200 µl of this working reagent to each well. 4. Cover the plate with foil and incubate for 30 min at 37˚C. 5. Read the plate at an absorbance of 562 nm.

1.5.4 IDENTIFICATION

OF

APP

AND ITS

DERIVATIVES

BY

WESTERN BLOT

1. Lyse samples with 3× sample buffer (10% SDS, 0.3 M Tris, 50% glycerol, 10% β-mercaptoethanol and a trace amount of bromophenol blue). 2. Heat samples at 100˚C for 5 min. 3. For BioRad Criterion gels, run at 200 V in running buffer (25 mM Tris, 192 mM glycine, 1% (w/v) SDS, pH 8.3) for 50 to 55 min. 4. Transfer gel to a 0.2-µm supported nitrocellulose membrane (BioRad, #162-0097) at 100 V for 1 hr at 4˚C in transfer buffer (20% methanol, 25 mM Tris, 192 mM glycine, 1% (w/v) SDS, pH 8.3). 5. Block the membrane in 5% milk in PBS-T (0.05% Tween-20) with agitation for 30 min at room temperature. 6. Wash two times with PBS-T for 2 min. 7. Incubate in primary antibody in PBS-T overnight at 4˚C or for 2 hr at room temperature. 8. Wash for 15 min, then wash for 5 min, three times in PBS-T. 9. Incubate 1 hr at room temperature in secondary antibody diluted 1:10,000 in PBS-T. The type of secondary antibody (antimouse, antirabbit, etc.) will depend on the primary antibodies used (monoclonal/mouse vs. polyclonal/rabbit antibodies). 10. Wash for 15 min, then wash for 5 min, three times in PBS-T. 11. Place the membrane on a transparency and add ECL Plus mixture (1 ml A plus 25 µl B; Amersham) to cover entire membrane for 1 min. 12. Place second transparency over membrane and expose for various periods (e.g., 15 sec, 30 sec, 1 min, and 5 min).

1.5.5 RADIOLABELING

OF

CELLS

WITH [35S]-MET

1. Aspirate media from cells cultured in 35-, 60-, or 100-mm dishes. Incubate cells with methionine-free medium for 15 to 30 min at 37˚C. 2. Add appropriate volume of methionine-free medium (0.75 ml for 35-mm dish, 2 ml for 60-mm dish and 5 ml for 100-mm dish). 3. Add [35S]-methionine to each dish to reach a final specificity of 100 to 200 µCi/ml. Incubate at 37˚C for the desired time; for abundant proteins, 2 to 4 hr is generally sufficient. If labeling for >6 hr, it may be necessary to supplement with fetal calf serum (10% or less) or 5 to 10% DMEM. 4. Collect cells for immunoprecipitation of target proteins. For secreted proteins, collect medium for immunoprecipitation.

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1.5.6 IDENTIFICATION OF FULL-LENGTH APP AND ITS DERIVATIVES BY IMMUNOPRECIPITATION 1. Culture cells to confluence in a 10-cm dish (~3 mg of protein), then briefly wash them twice in PBS. (For storage, 2 ml of 20 mM EDTA in PBS is added and cells are collected and centrifuged at 3500 g for 5 min. Cell pellets can be frozen at –80˚C.) 2. Cells are lysed in an IP Lysis buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 2 mM EDTA, 1% NP-40 and a protease inhibitor cocktail [5 µg/ml leupeptin, 5 µg/ml aprotinin, 2 µg/ml pepstatin A, and 0.25 mM phenylmethylsulfonyl fluoride (Sigma)]). Incubate lysates on ice for 20 min. Centrifuge lysates at 3500 g for 5 min. Transfer supernatant to a fresh tube. 3. Lysates are precleared with 20 µl of protein A-sepharose CL-4B (Sigma, #P-3391) at 100 mg/ml in STEN buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 2 mM EDTA, 0.2% NP-40) for 0.5 hr at 4˚C. Supernatants are transferred for immunoprecipitation with primary antibodies with 20 µl protein A-sepharose or protein G-agarose (for a monoclonal antibody) at 4˚C for 2 hr. 4. Immunoprecipitates are washed in a 0.5 M STEN buffer (50 mM Tris, pH 7.6, 500 mM NaCl, 2 mM EDTA, and the same protease inhibitor cocktail described above) for 15 min at 4˚C. Centrifuge the samples at 3500 g for 5 min at 4˚C and discard the supernatant. 5. Immunoprecipitates are washed in a SDS-STEN buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 0.1% SDS, 2 mM EDTA, and the protease inhibitor cocktail) for 15 min at 4˚C. Centrifuge the samples at 3500 g for 5 min at 4˚C and discard the supernatant. 6. Immunoprecipitates are washed in a STEN buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 0.2% NP-40, 2 mM EDTA, and the protease inhibitor cocktail) for 15 min at 4˚C. Centrifuge the samples at 3500 g for 5 min at 4˚C. Discard the supernatant, then elute the immunoprecipitates with 3× sample buffer (10% SDS, 0.3 M Tris, 50% glycerol, 10% β-mercaptoethanol and a trace amount of bromophenol blue), heat at 100˚C for 5 min, and separate the samples on a 4 to 20% tris-glycine gel by SDS-PAGE (BioRad, Criterion). 7. Gels are stained in Coomassie blue [0.2% Coomassie in destain solution (50% methanol, 18% acetic acid)] followed by destaining in destain solution for 30 to 45 min to fix proteins. Dry gel after washing gel in Gel-Dry solution (Invitrogen, #LC4025-4) for 30 min. Expose gel in phosphorimager for imaging or to film at –80˚C. 8. Alternatively, confluent cells can be directly lysed in lysis buffer without radiolabeling, and immunoprecipitates can be separated by electrophoresis followed by Western blot, using standard procedures provided by the manufacturer, e.g., ECL Plus detection kit from Amersham.

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1.5.7 DETERMINATION

OF

HALF-LIFE

OF

APP

The half-life of a target protein can be determined by pulse-chase labeling of cells followed by immunoprecipitation with its specific antibody. In addition to determining the rate of holoprotein turnover, the course of protein maturation can also be observed. 1. Growth media of confluent CHO cells are replaced with methionine (Met)free media, and cells are incubated at 37˚C for 45 min before the media are aspirated. 2. Cells are pulse labeled with prewarmed Met-free media containing 100 µCi/ml of [35S]-Met for 5 to 15 min. 3. [35S]-Met-containing media are removed, and cells are washed twice with regular DMEM media. 4. Cells are chased in prewarmed DMEM media for an appropriate period before they are collected for immunoprecipitation.

1.5.8 CO-IMMUNOPRECIPITATION

OF

APP-INTERACTING PROTEIN

1.5.8.1 Preparation 1. Thaw Co-IP lysis buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 2 mM EDTA, 1% NP-40, 0.5% Triton X-100, 0.5% BSA, and a protease inhibitor cocktail) at room temperature. 2. Thaw cell pellets/membrane vesicles on ice, and leave protein A-sepharose on ice. 1.5.8.2 Pre-Absorption of Protein A-Sepharose 1. Pipet 2 ml of Co-IP lysis buffer into an Eppendorf tube, then add up to 200 µl of protein-A sepharose. Since each sample will be equally divided and co-immunoprecipitated with an immune antibody or a control preimmune serum, 40 µl of protein A-sepharose will be needed for each sample of cell lysate. 2. Rotate the Eppendorf tube for at least 4 hr at 4˚C (in the cold room). 1.5.8.3 Pre-Clearing of Cell Lysates 1. Lyse the pellets/vesicles with 1 ml of Co-IP lysis buffer and incubate on ice for 20 min. 2. Centrifuge at 3500 g for 5 min. 3. Transfer supernatant into a new tube. 4. Vortex protein A-sepharose briefly, and add 20 µl of protein A-sepharose into each tube. 5. Rotate samples for at least 4 hr at 4˚C (in the cold room).

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1.5.8.4 Set Up Co-IP 1. Label new Eppendorf tubes: two per cell lysate, one for the sample to be immunoprecipitated with preimmune serum (Pre) and the other for the immune serum. 2. Pipet 2 µl of preimmune serum (1:200 dilution) into each preimmune sample. For easier pipetting, create a larger volume by combining the total amount of preimmune serum for all Pre samples with about 100 µl of lysis buffer. Add equal amounts to each tube. 3. For a co-IP of presenilin, for example, put 1 µl of antibody X81 and 1 µl of antibody 4627 (1:200 dilution; dilution ratio should be the same as that of preimmune; adjust if necessary) to each sample. For easier pipetting, add the total amounts of X81 and 4627 for all immune samples into a tube with about 100 µl of lysis buffer. Add equal amounts to each tube. 4. Spin down the cell samples and preabsorbed protein A-sepharose at 3500 g for 5 min. 5. Equally divide the supernatant from each sample and aliquot it into two tubes that contain preimmune serum or immune serum. 6. Remove most of the supernatant of preabsorbed protein A-sepharose, and leave a sufficient amount of buffer equivalent to two times the volume of protein A-sepharose. 7. Vortex preabsorbed protein A-sepharose and transfer 20 µl into each tube. 8. Rotate samples for at least 4 hr (in the cold room). 1.5.8.5 Wash Co-IP Two solutions will be used: 0.5 M STEN (50 mM Tris, pH 7.6, 500 mM NaCl, 2 mM EDTA, and the protease inhibitor cocktail) and STEN (50 mM Tris, pH 7.6, 150 mM NaCl, 0.2% NP-40, 2 mM EDTA, and the protease inhibitor cocktail). 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Spin samples at 3500 g for 5 min. Aspirate the supernatant without touching the immunoprecipitates. Pipet 750 µl of 0.5 M STEN into each tube. Rotate samples for 15 min (in the cold room). Spin samples at 3500 g for 5 min. Aspirate the supernatant. Pipet 750 µl STEN into each tube. Rotate samples for 15 min in the cold room. Spin samples at 3500 g for 5 min. Aspirate most, but not all, of the supernatant. Use a P20 Pipetman to remove the last of the supernatant. Do not take up any beads. 11. Add 20 µl of 3× sample buffer (10% SDS, 0.3 M Tris, 50% glycerol, 10% β-mercaptoethanol, and a trace amount of bromophenol blue) into each sample.

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12. 13. 14. 15.

Vortex the samples, and heat at 100˚C for 5 min. Spin at 18,000 g for 5 min. Load samples into the gel. Do not take up any beads. Proceed to Western blot for detection of co-immunoprecipitates.

Many elements must be monitored carefully for successful detection of protein complexes. Several key components are: 1. Proper preabsorption of protein A-sepharose and preclearing of cell lysate will reduce nonspecific binding of proteins to protein A-sepharose. Without these two steps, the complex will be immunoprecipitated in samples incubated with preimmune serum due to nonspecific binding to protein A-sepharose. 2. The choice of detergents affects the stringency of co-immunoprecipitation as well as the maintenance of the intact complex. Another detergent that can be used to replace 0.5% Triton X-100 and 1% NP-40 is 1% CHAPSO. A battery of detergents must be tested to detect a specific interaction between two proteins.

1.5.9 CONJUGATION

OF

ANTIBODY

TO

PROTEIN A-SEPHAROSE

Most co-immunoprecipitation experiments are carried out using unconjugated antibody (e.g., serum) and protein A-sepharose (or protein G-agarose), and the immunoprecipitates are detected by Western blot. The best method is to use polyclonal antibodies for immunoprecipitation and monoclonal antibodies for Western blot. However, if the choice of antibodies is limited and the same type of antibody must be used for both immunoprecipitation and Western blot, the cross-reactivity of IgG heavy and light chains eluted from the immunoprecipitates will lead to a much higher background at molecular weights of ~25 kDa and above ~55 kDa. Although it does not completely eliminate free heavy and light chains, conjugation of primary antibody for immunoprecipitation to protein A-sepharose will significantly reduce the number of IgG heavy and light chains eluted from immunoprecipitates, thus reducing the background of the Western blot. 1. Suspend 2 g of protein A-sepharose beads in 16 ml of conjugation buffer (50 mM Tris, pH 7.6, 150 mM NaCl, 2% BSA) to make a final concentration of 125 mg/ml protein A-sepharose. This will give a final yield of eight tubes of 2-ml conjugated beads. 2. Combine each 1 ml of resuspended beads with 125 µl antiserum. Incubate 1 hr at room temperature with rocking, then place 2 ml of beads plus antiserum per 15 ml conical tube. 3. Centrifuge for 5 min at 3000 g. Save the supernatant in case of a coupling problem. 4. Resuspend the beads in 10 ml of 0.2 M sodium borate buffer, pH 9.0 (heat to dissolve the precipitates prior to usage). Centrifuge the beads and repeat the wash as before. Bring up in 10 ml of sodium borate buffer. Remove 100 µl of supernatant to check later (A).

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5. Add 50 mg dimethyl pimelimidate (DMP; Sigma, #D8388, store at –20˚C) to each tube. Ensure that pH >8.3. Incubate for 30 min at room temperature with rocking. Remove 100 µl of supernatant to check later (B). 6. To stop the reaction, spin the beads at 3000 g for 5 min. Add 10 ml 0.2 M ethanolamine (pH 8.0) per tube. Wash one more time with 10 ml 0.2 M ethanolamine, then incubate in 10 ml ethanolamine for 2 hr at room temperature with rocking. 7. Centrifuge and transfer each pellet to a 2-ml Eppendorf tube. 8. Add 1 ml 100 mM glycine, pH 3.0; mix and then spin at 10,000 g for 30 sec. Remove the supernatant and wash one time with 1 ml 100 mM Tris pH 8.0; mix and spin. Expose beads to glycine buffer for as short a time as possible. 9. Resuspend beads in 1 ml PBS with 0.01% thimerosal for each tube. Remove 100 µl of supernatant to check later (C). To check the efficiency of conjugation, the amount of remaining IgG can be detected in the supernatant (A, B, and C) by Western blot. With the above protocol, the conjugated antibodies with protein A-sepharose are usually stable for >1 year at 4˚C.

ACKNOWLEDGMENTS This work was supported in part by National Institutes of Health (AG 17593) and the Foundation for Neurologic Diseases. I would like to thank many of my colleagues at the Center for Neurologic Diseases, especially Dr. Dennis Selkoe, who fosters a superb research environment.

REFERENCES 1. Selkoe, D.J. The genetics and molecular pathology of Alzheimer’s disease, in Neurologic Clinics: Dementia, DeKosky, S.T. et al., Eds., W.B. Saunders, Philadelphia, 2000, 18, 903–921. 2. Selkoe, D.J. and Podlisny, M.B. Deciphering the genetic basis of Alzheimer’s disease. Annu. Rev. Genomics Hum. Genet. 3, 67–99, 2002. 3. Selkoe, D.J. Cell biology of the amyloid β-protein precursor and the mechanism of Alzheimer’s disease. Annu. Rev. Cell Biol. 10, 373–403, 1994. 4. Kang, J. et al. The precursor of Alzheimer’s disease amyloid A4 protein resembles a cell-surface receptor. Nature 325, 733–736, 1987. 5. Goate, A. et al. Segregation of a missense mutation in the amyloid precursor protein gene with familial Alzheimer’s disease. Nature 349, 704–706, 1991. 6. Hendriks, L. et al. Presenile dementia and cerebral haemorrhage linked to a mutation at codon 692 of the β-amyloid precursor protein gene. Nature Genet. 1, 218–221, 1992. 7. Levy, E. et al. Mutation of the Alzheimer’s disease amyloid gene in hereditary cerebral hemorrhage, Dutch-type. Science 248, 1124–1126, 1990. 8. van Broeckhoven, C. et al. Amyloid β-protein precursor gene and hereditary cerebral hemorrhage with amyloidosis (Dutch). Science 248, 1120–1122, 1990.

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9. Mullan, M. et al. A pathogenic mutation for probable Alzheimer’s disease in the APP gene at the N-terminus of β-amyloid. Nature Genet. 1, 345–347, 1992. 10. Eckman, C. et al. A new pathogenic mutation in the APP gene (I716V) increases the relative proportion of Aβ 42(43). Human Mol. Genet. 6, 2087–2089, 1997. 11. Chartier-Harlin, M.C., Crawford, F., and Houlden, H. Early-onset Alzheimer’s disease caused by mutations at codon 717 of the β-amyloid precursor protein gene. Nature 353, 844–846, 1991. 12. Murrell, J., Farlow, M., Ghetti, B., and Benson, M.D. A mutation in the amyloid precursor protein associated with hereditary Alzheimer’s disease. Science 254, 97–99, 1991. 13. Neve, R.L., Finch, E.A., and Dawes, L.R. Expression of the Alzheimer amyloid precursor gene transcripts in the human brain. Neuron 1, 669–677, 1988. 14. Citron, M. et al. Mutation of the β-amyloid precursor protein in familial Alzheimer’s disease increases β-protein production. Nature 360, 672–674, 1992. 15. Cai, X.D., Golde, T.E., and Younkin, G.S. Release of excess amyloid β protein from a mutant amyloid β protein precursor. Science 259, 514–516, 1993. 16. Citron, M. et al. Excessive production of amyloid β-protein by peripheral cells of symptomatic and presymptomatic patients carrying the Swedish familial Alzheimer’s disease mutation. Proc. Natl. Acad. Sci. USA 91, 11993–11997, 1994. 17. Scheuner, D. et al. Secreted amyloid β-protein similar to that in the senile plaques of Alzheimer’s disease is increased in vivo by the presenilin 1 and 2 and APP mutations linked to familial Alzheimer’s disease. Nature Med. 2, 864–870, 1996. 18. Suzuki, N. et al. An increased percentage of long amyloid β protein secreted by familial amyloid β protein precursor (βAPP717) mutants. Science 264, 1336–1340, 1994. 19. Games, D. et al. Alzheimer-type neuropathology in transgenic mice overexpressing V717F β-amyloid precursor protein. Nature 373, 523–527, 1995. 20. Hsiao, K. et al. Correlative memory deficits, Aβ elevation, and amyloid plaques in transgenic mice. Science 274, 99–102, 1996. 21. Vassar, R. et al. β-secretase cleavage of Alzheimer’s amyloid precursor protein by the transmembrane aspartic protease BACE. Science 286, 735–741, 1999. 22. Sinha, S. et al. Purification and cloning of amyloid precursor protein β-secretase from human brain. Nature 402, 537–540, 1999. 23. Yan, R. et al. Membrane-anchored aspartyl protease with Alzheimer’s disease β-secretase activity. Nature 402, 533–537, 1999. 24. Lin, X. et al. Human aspartic protease memapsin 2 cleaves the β-secretase site of β-amyloid precursor protein. Proc. Natl. Acad. Sci. USA 97, 1456–1460, 2000. 25. Seubert, P. et al. Secretion of β-amyloid precursor protein cleaved at the aminoterminus of the β-amyloid peptide. Nature 361, 260–263, 1993. 26. Haass, C. et al. Amyloid β-peptide is produced by cultured cells during normal metabolism. Nature 359, 322–325, 1992. 27. Shoji, M. et al. Production of the Alzheimer amyloid β protein by normal proteolytic processing. Science 258, 126-129, 1992. 28. Lammich, S. et al. Constitutive and regulated α-secretase cleavage of Alzheimer’s amyloid precursor protein by a disintegrin metalloprotease. Proc. Natl. Acad. Sci. USA 96, 3922–3927, 1999. 29. Buxbaum, J.D. et al. Evidence that tumor necrosis factor alpha converting enzyme is involved in regulated α-secretase cleavage of the Alzheimer amyloid protein precursor. J. Biol. Chem. 273, 27765–27767, 1998.

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30. Esch, F.S. et al. Cleavage of amyloid β-peptide during constitutive processing of its precursor. Science 248, 1122–1124, 1990. 31. Sisodia, S.S., Koo, E.H., Beyreuther, K., Unterbeck, A., and Price, D.L. Evidence that β-amyloid protein in Alzheimer’s disease is not derived by normal processing. Science 248, 492–495, 1990. 32. Haass, C., Hung, A.Y., Schlossmacher, M.G., Teplow, D.B., and Selkoe, D.J. β-Amyloid peptide and a 3-kDa fragment are derived by distinct cellular mechanisms. J. Biol. Chem. 268, 3021–3024, 1993. 33. Bretz, R. and Staubli, W. Detergent influence on rat-liver galactosyl transferase activities towards different acceptors. Eur. J. Biochem. 77, 181–192, 1977. 34. Weidemann, A. et al. Identification, biogenesis and localization of precursors of Alzheimer’s disease A4 amyloid protein. Cell 57, 115–126, 1989. 35. Oltersdorf, T. et al. The Alzheimer amyloid precursor protein: identification of a stable intermediate in the biosynthetic/degradative pathway. J. Biol. Chem. 265, 4492–4497, 1990. 36. Haass, C. et al. Swedish mutation causes early-onset AD by β-secretase cleavage within the secretory pathway. Nature Med. 1, 1291–1296, 1995.

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2

Assays for Analysis of APP Secretion and Recycling Markus P. Kummer, Tsuneo Yamazaki, and Edward H. Koo

CONTENTS Abstract 2.1 Introduction 2.2 Main Scheme of Approaches 2.3 Methods 2.3.1 Iodination of Antibody 2.3.2 Preparation of Cells 2.3.3 Kinetics of Secretion and Endocytosis of APP 2.3.4 Steady State Level of APP Endocytosis 2.3.5 Recycling of APP 2.3.6 Morphological Analysis 2.4 Discussion Acknowledgments References

ABSTRACT The trafficking of the amyloid precursor protein (APP) involves the concomitant secretion and endocytosis of APP from the cell surface. In addition, APP recycles between the endocytic compartment and the cell surface. This complex sequence of events can be studied by a reliable and reproducible assay based on the binding of a radiolabeled APP antibody. Using this technique, the secretion and internalization of APP can be measured simultaneously under normal and perturbated conditions in APP transfected cells. Furthermore, this method is readily adaptable to morphologically examine the pathways of APP trafficking from the cell surface.

0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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2.1 INTRODUCTION APP is a type I membrane protein that undergoes constitutive shedding both intracellularly and at the cell surface to release a large secreted ectodomain derivative (sAPP). APP is transported in the secretory pathway from the golgi to the plasma membrane where it can be cleaved by α-secretase to release the N terminal APP ectodomain (sAPPα) or internalized and transported to the endosomal–lysosomal pathway. APP trafficking has been actively investigated in order to determine the cellular sites where Aβ is generated. From these studies, a major pathway of Aβ generation is from APP that is processed following internalization from the cell surface in the endocytic pathway, although the precise organelles where β- and γ-secretase cleavages take place remain to be defined.1 APP internalization is mediated via clathrin-coated pits2,3 in the canonical receptor-mediated endocytic pathway. Recently, however, it has been shown that a fraction of APP is associated with detergent-resistant membranes and that lipid rafts are apparently sites of Aβ production as well.4–7 Whether raft-associated APP is internalized and, if so, whether this pool subsequently merges with the nonraft pool is unclear. Following internalization, a fraction of APP returns to the cell surface for additional rounds of endocytosis or secretion. The internalization signal for APP resides within its cytoplasmic domain and belongs to the NPXY-type signal, one of several well-recognized motifs. This signal is present in APP, β-integrin, and low density lipoprotein receptor (LDLR) protein family members among others, and controls the rapid internalization of these integral membrane proteins.8 Although this signal represents the minimal amino acid sequence shared by all these proteins, it may be not sufficient for efficient internalization. In LDLR, an additional aromatic amino acid residue upstream of the NPXY motive has become an accepted signal for internalization. In the case of APP, the signal is mediated by the longer GYENPTY sequence. Mutagenesis studies of this motif revealed that the predominant signal lies in the YENP tetrapeptide.9 Several cytosolic adaptor proteins such as Fe65, X11, and the mammalian Dab1 bind to the cytoplasmic YENPTY motif via their phosphotyrosine interaction domains.10–12 They affect the subcellular trafficking and proteolytic processing of APP in different ways. Fe65, for example, increases the secretion of APP and promotes A secretion, whereas X11 retards APP catabolism and inhibits Aβ secretion.13,14

2.2 MAIN SCHEME OF APPROACHES The analysis of APP processing in the endocytic pathway has been difficult because of the concurrent secretion and internalization of APP molecules from the cell surface. The assay described presents a reliable and reproducible method to measure the secretion of sAPP and APP internalization from the cell surface simultaneously. Radiolabeled antibodies have been successfully used as surrogates to natural ligands to investigate the internalization of other transmembrane receptors like LDLR, transferrin, CD4, or macrophage Fc receptors.15–18 This protocol contains the procedure for radioiodination of the monoclonal APP antibody, 1G7, and four variations for analyzing the trafficking of APP (Figure 2.1).

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Incubation with radiolabeled antibody

at 37°C

Steady-state level of internalization (method 2.3.4)

at 4°C

Labeling of cell surface pool Wash and incubate at 37°C

Cool to 4°C, acid wash, warm to 37°C

Recycled molecules (method 2.3.5)

Kinetics of secretion and internalization (method 2.3.3)

FIGURE 2.1 Schematical overview about the different methods to characterize APP trafficking using a radiolabeled antibody.

The first method describes a pulse/chase experiment to analyze the kinetics of APP secretion and internalization. In the second method, APP internalization is investigated under steady state conditions. The third method addresses the recycling of APP to the cell surface. The fourth and final is a morphological approach to examine APP internalization. These methods using the monoclonal antibody 1G7 have been successfully applied to determine the trafficking of APP in a variety of cells including primary neurons and transfected CHO, B103, and N2a cell lines.9,19–21

2.3 METHODS 2.3.1 IODINATION

OF

ANTIBODY

The monoclonal antibody 1G7 was raised against human APPs purified from APPtransfected CHO cells and recognizes an epitope in the extracellular domain of APP between residues 380 and 665, as defined by its reactivity against a bacterial fusion protein with this sequence, thereby excluding both the KPI and Aβ domains.19 The specificity of this antibody for APP was demonstrated previously by immunoprecipitation and immunofluoresence studies.1,22 The 1G7 antibody is radioiodinated using IODO-GEN precoated iodination tubes (Pierce, Rockford, IL; #28601). This method results in indirect labeling without contact of the antibody to the iodination reagent, thereby reducing oxidative damage

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to the antibody. Excessive iodine is finally removed by gel filtration, typically with a disposable NAP-5 (Sephadex G-25) column (Amersham Biosciences, Piscataway, NJ; #17-0853-01). The entire procedure must be performed in an approved protective hood to avoid uptake of free radioactive iodine. 1. Pre-equilibrate a NAP-5 column with 10 ml of Dulbecco’s phosphatebuffered saline (DPBS). 2. Rinse an IODO-GEN-coated iodination tube with 0.5 ml DPBS. 3. Add 300 µl DPBS directly to the bottom of the tube. Add 1 mCi Na125I in 10 mM NaOH. Incubate for 6 min, gently swirling the tube every 30 sec. This step results in the generation of iodous ions (I+) by oxidation of iodide (I–) by the iodination reagent. 4. Remove and add the activated iodide to 200 µl DPBS containing 100 µg of 1G7 antibody in a new screw cap tube. The final volume with the activated iodide solution is 500 µl. 5. Incubate the mixture for 6 min, gently flicking the tube every 30 sec. During this incubation, iodous ions undergo electrophilic attack at the ortho ring positions of tyrosine residues. 6. Remove the labeling solution and add it to a pre-equilibrated NAP-5 column. Drain the column and discard the flow-through. 7. Elute the column with 980 µl DPBS. Add 20 µl of 100 mg/ml crystalline BSA as a carrier protein to a final concentration of 2 mg/ml. 8. Determine radiospecific activity of the labeled antibody in a gamma counter. Typically, the radiospecific activity will be 7,500 to 15,000 cpm/fmol (3 to 6 µCi/µg) when 66 nm (100 µg) of antibody is labeled by this procedure. In our experience, the antibody is stable for up to 4 weeks when stored at 4˚C.

2.3.2 PREPARATION

OF

CELLS

APP-transfected cells are seeded in 12 well plates 48 hr before the assay and grown to confluence. Nonspecific binding by the antibody and the endogenous APP signal are subtracted by analyzing untransfected cells grown in parallel. Alternatively, radiolabeled antibody binding can be competed with excess cold antibody.

2.3.3 KINETICS

OF

SECRETION

AND

ENDOCYTOSIS

OF

APP

This protocol examines the trafficking of APP by pulse/chase analysis. A population of surface APP molecules is initially bound to radiolabeled antibody at 4˚C and is then, after removal of unbound antibody, allowed to transit when rewarmed to 37˚C. At various time points, the supernatant containing the secreted APP is collected and any remaining antibody is removed from the cell surface by acid washes. Finally, the cell pellet is lysed (acid resistant fraction) to obtain the internalized APP fraction and all samples are measured in a gamma counter.

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1. Chill the cells on ice to stop membrane trafficking. 2. Wash the cells twice with 2 ml of ice cold binding medium consisting of RPMI containing 20 mM HEPES (pH 7.5) supplemented with 0.2% bovine serum albumin. 3. Add approximately 0.4 µg 125I-IG7 antibody in 400 µl binding medium per well (~7 nM final) and allow binding of the antibody for 1 hr on ice. The Kd for this reaction is 1.23 nM for a CHO cell line stably overexpressing APP.19 Therefore, this concentration is at least fivefold the concentration needed for half-maximal saturation. 4. Wash the cells twice with binding medium and twice with ice cold DPBS. 5. Place the cells in 2 ml of prewarmed medium and incubate at 37˚C for various periods. 6. Collect the medium and chill cells rapidly by adding 2 ml of ice cold DPBS at pH 2.5. Incubate the cells for another 5 min. Collect the supernatant, repeat the acidic wash to remove residual surface-bound antibody and pool both supernatants. The acid wash detaches 90 to 95% of cell surface-bound antibody 7. Lyse the cells by adding 2 ml of 0.2 N NaOH. 8. Measure the radioactivity of all fractions in a gamma counter. To calculate the specific binding, the radioactivity from untransfected control cells is subtracted from the counts obtained from the transfected cells in each condition. The results obtained at each time point are expressed as a percentage of total radioactivity from the three fractions (medium, acid wash and cell lysate). The anticipated result should show a rapid release of sAPP into the medium with a half life of 10 min. Consequently, cell surface APP recovered by the two acid washes should rapidly decline and remain at low levels. The remaining cell surface APP is internalized within 10 min of rewarming to 37˚C. The internalized pool declines concurrently with an increase in the secreted pool, because of recycling of APP to the cell surface and subsequent secretion into the medium. After 30 min, the secreted pool remains stable whereas the internalized fraction declines, presumably because of degradation. If the experiment is prolonged to more than 30 min, one should take into account that some of the radiolabeled antibody may be degraded and therefore free radioactivity-liberated. Under these circumstances, the medium and cell lysate should first be precipitated with trichloroacetic acid to recover the antibody-bound radioactivity only. As a consequence of the precipitation, the total radioactivity will be less than 100%.

2.3.4 STEADY STATE LEVEL

OF

APP ENDOCYTOSIS

To measure the rate of APP endocytosis under steady state conditions, the radiolabeled antibody is allowed to bind at 37˚C, resulting in a concomitant uptake of radioiodinated antibody by APP internalization. This follows the method established for assessing

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the internalization of transferrin receptor under steady state conditions. After incubation, the cell surface-bound pool is removed by acid washes whereas the internalized fraction is recovered by cell lysis. This straightforward method is a facile approach for comparing the efficiency of APP endocytosis in cell lines transfected with different APP mutants or constructs of other proteins that may affect APP internalization. 1. Wash the cells twice with prewarmed binding medium. 2. Add approximately 7 nmol 125I-IG7 antibody in 400 µl binding medium per well and allow binding and internalization of the antibody for 30 to 60 min at 37˚C. 3. Chill the cells on ice and wash the cells four times with ice cold DPBS. 4. Remove the cell surface-bound antibody by two consecutive washes with 2 ml of ice cold DPBS at pH 2.5 for 5 min each on a shaker and pool both supernatants. 5. Lyse the cells by adding 2 ml of 0.2 N NaOH. 6. Measure the radioactivity of both fractions in a gamma counter. The radioactivity from untransfected control cells is subtracted from the counts resulting from the transfected cells to obtain specific binding values. The results are expressed as percents of the cell lysate (acid-resistant) and cell surface (acid-labile) fractions to reflect the percent of APP internalized from the cell surface.

2.3.5 RECYCLING

OF

APP

Internalized APP recycles and is secreted 10 to 30 min after endocytosis.19 The amounts of APP and secreted APP released from the endocytosed pool are analyzed by incubation of the cells with radioiodinated antibody at 37˚C and subsequent removal of any cell surface-bound antibody by acid washes. After rewarming the medium containing secreted APP is collected and cell surface APP is detached by two consecutive acid washes. The medium is precipitated with trichloroacetic acid to recover the antibody-bound radioactivity only. 1. Wash the cells twice with prewarmed binding medium. 2. Add approximately 7 nmol 125I-IG7 antibody in 400 µl binding medium per well and allow binding and internalization of the antibody for 15 min at 37˚C. 3. Wash the cells four times with ice cold DPBS. 4. Remove the cell surface-bound antibody by two consecutive washes with 2 ml of ice cold DPBS at pH 2.5 for 3 min each and discard the supernatants. 5. Wash the cells twice with ice cold binding medium, then add prewarmed medium and return cells to the 37˚C incubator. 6. Collect the medium after 5 to 30 min. Precipitate the medium with TCA. 7. Lyse the cells by adding 2 ml of 0.2 N NaOH. 8. Measure the radioactivity of both fractions in a gamma counter.

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To obtain the specific binding, the radioactivity from untransfected control cells is subtracted from the counts resulting from the transfected cells. The radioactivity from the medium is computed after precipitation with trichloroacetic acid to measure the antibody-bound radioactivity. The results are expressed as radioactivity from the TCA precipitation as a percentage of the radioactivity from both fractions (lysate and TCA precipitate). The expected result should be a gradual increase of APP in the medium over 30 min, suggesting that internalized APP and therefore the bound radiolabeled antibody are recycled to the cell surface and secreted into the medium. In contrast to that, the counts in the pellets should decrease.

2.3.6 MORPHOLOGICAL ANALYSIS To visualize directly the trafficking routes of cell surface APP, immunofluorescence detection of APP antibody added to living cells combined with suitable organelle markers is a straightforward method to morphologically evaluate the internalization pathways of APP localization sites. However, it is important to be aware that the antibody may induce perturbations to the normal internalization pathways. To avoid cross-linking of cell surface APP, the results from whole antibody should be confirmed with Fab fragments or with other antibodies recognizing different epitopes. The choice of cell types is also an important consideration. The ideal cells should be adherent and have large cytoplasms for easy visualization. Importantly, the cells must remain adherent during the many washes and incubations on ice. Rat hippocampal neurons cultured at low density are also suitable, but the neuritic processes are easily damaged during the procedure. The following is our standard protocol for visualizing internalization of cell surface APP in CHO cells. 1. CHO cells stably transfected with APP 751 are grown on coverslips and cultured in standard medium. 2. The cells are chilled on ice for 15 min and washed with ice cold DPBS to stop membrane trafficking. 3. Coverslips are then incubated with anti-APP monoclonal antibodies (1G7 alone or combined with 5A3) in cold DPBS for 1 hr on ice. 4. The cells are washed 5 times with cold DPBS and then incubated with prewarmed regular medium for various times (0 to 60 min) at 37°C. The cells are then fixed with cold 4% formaldehyde (freshly prepared from paraformaldehyde) in PBS for 15 min. 5. Following fixation, the cells are permeabilized for 5 min with 0.3% Triton X-100 in PBS, washed 3 times with PBS, and incubated with fluorescein isothionate (FITC)-conjugated anti-mouse secondary antibody for 1 hr at room temperature. 6. If desired, double labeling with another antibody, such as to organelle marker, can be used at this point. 7. The cover slips are mounted on slide glasses and visualized with conventional epifluorescence or confocal microscopy.

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2.4 DISCUSSION In this chapter, we have described assays based on the binding of a radioiodinated or unlabeled monoclonal antibody to investigate the aspects of APP trafficking in cultured cells. The use of an antibody for kinetic analysis of APP trafficking from the cell surface may be the only possible approach because of the lack of a physiological soluble ligand for APP at the cell surface. Biotinylation of cell surface APP is possible but this would be very cumbersome for obtaining kinetic information. The 1G7 antibody used in this protocol has been shown to bind to cell surface APP in a saturable and competitive manner. This antibody is also not labile at 37°C, i.e., not readily detached from APP. Other APP antibodies recognizing the extracellular region of APP can certainly be used, but these same parameters should be tested first. Additionally, the concentration for half-maximal saturation should be measured by Scatchard plot analysis to determine the antibody concentration necessary for the assay. To verify that the experimental conditions do not result in a general perturbation of endocytosis, the internalization of another transmembrane protein should be measured. In this case, the uptake of transferrin by the transferrin receptor is frequently used as a control.21 For this purpose bovine holo-transferrin (Sigma) is iodinated as described and added to the cells exactly as described by Zuk et al.23 In summary, the approach described in this chapter provides rapid and accurate estimates for APP secretion and endocytosis as well as morphological assessment of the internalization pathways. In this way, the complex APP trafficking pathways in neurons and non-neural cells can be analyzed. In particular, the influence of mutations within the cytoplasmic domains of APP or the impacts of cytosolic APP binding proteins on the processing of APP can be determined.

ACKNOWLEDGMENTS This work was supported in part by NIH Grant AG 12376. We thank Drs. Christian Haass, Claus Pietrzik, Ruth Perez, and Dennis Selkoe for helpful discussions.

REFERENCES 1. Koo, E.H. and Squazzo, S.L. Evidence that production and release of amyloid betaprotein involves the endocytic pathway. J. Biol. Chem. 269, 17386, 1994. 2. Nordstedt, C. et al. Identification of the Alzheimer beta/A4 amyloid precursor protein in clathrin-coated vesicles purified from PC12 cells. J. Biol. Chem. 268, 608, 1993. 3. Yamazaki, T., Koo, E.H., and Selkoe, D.J. Trafficking of cell-surface amyloid betaprotein precursor. II. Endocytosis, recycling and lysosomal targeting detected by immunolocalization. J. Cell Sci. 109, 999, 1996. 4. Simons, M. et al. Cholesterol depletion inhibits the generation of beta-amyloid in hippocampal neurons. Proc. Natl. Acad. Sci. USA 95, 6460, 1998. 5. Aplin, A.E. et al. Effect of increased glycogen synthase kinase-3 activity upon the maturation of the amyloid precursor protein in transfected cells. Neuroreport 8, 639, 1997.

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6. Bouillot, C. et al. Axonal amyloid precursor protein expressed by neurons in vitro is present in a membrane fraction with caveolae-like properties. J. Biol. Chem. 271, 7640, 1996. 7. Ehehalt, R. et al. Amyloidogenic processing of the Alzheimer beta-amyloid precursor protein depends on lipid rafts. J. Cell Biol. 160, 113, 2003. 8. Bonifacino, J.S. and Traub, L.M. Signals for sorting of transmembrane proteins to endosomes and lysosomes. Annu. Rev. Biochem. 72, 395, 2003. 9. Perez, R.G. et al. Mutagenesis identifies new signals for beta-amyloid precursor protein endocytosis, turnover, and the generation of secreted fragments, including Abeta42. J. Biol. Chem. 274, 18851, 1999. 10. Borg, J.P. et al. The phosphotyrosine interaction domains of X11 and FE65 bind to distinct sites on the YENPTY motif of amyloid precursor protein. Mol. Cell. Biol. 16, 6229, 1996. 11. Fiore, F. et al. The regions of the Fe65 protein homologous to the phosphotyrosine interaction/phosphotyrosine binding domain of Shc bind the intracellular domain of the Alzheimer’s amyloid precursor protein. J. Biol. Chem. 270, 30853, 1995. 12. Howell, B.W. et al. The disabled 1 phosphotyrosine-binding domain binds to the internalization signals of transmembrane glycoproteins and to phospholipids. Mol. Cell. Biol. 19, 5179, 1999. 13. Borg, J.P. et al. The X11alpha protein slows cellular amyloid precursor protein processing and reduces Abeta40 and Abeta42 secretion. J. Biol. Chem. 273, 14761, 1998. 14. Sabo, S.L. et al. Regulation of beta-amyloid secretion by FE65, an amyloid protein precursor-binding protein. J. Biol. Chem. 274, 7952, 1999. 15. Pelchen-Matthews, A., Armes, J.E., and Marsh, M. Internalization and recycling of CD4 transfected into HeLa and NIH3T3 cells. EMBO J. 8, 3641, 1989. 16. Mellman, I.S. et al. Internalization and degradation of macrophage Fc receptors during receptor-mediated phagocytosis. J. Cell Biol. 96, 887, 1983. 17. Hopkins, C.R. and Trowbridge, I.S. Internalization and processing of transferrin and the transferrin receptor in human carcinoma A431 cells. J. Cell Biol. 97, 508, 1983. 18. Beisiegel, U. et al. Monoclonal antibodies to the low density lipoprotein receptor as probes for study of receptor-mediated endocytosis and the genetics of familial hypercholesterolemia. J. Biol. Chem. 256, 11923, 1981. 19. Koo, E.H. et al. Trafficking of cell-surface amyloid beta-protein precursor. I. Secretion, endocytosis and recycling as detected by labeled monoclonal antibody. J. Cell Sci. 109, 991, 1996. 20. Soriano, S. et al. Expression of beta-amyloid precursor protein-CD3gamma chimeras to demonstrate the selective generation of amyloid beta(1-40) and amyloid beta(142) peptides within secretory and endocytic compartments. J. Biol. Chem. 274, 32295, 1999. 21. Pietrzik, C.U. et al. The cytoplasmic domain of the LDL receptor-related protein regulates multiple steps in APP processing. EMBO J. 21, 5691, 2002. 22. Yamazaki, T., Selkoe, D.J., and Koo, E.H. Trafficking of cell surface beta-amyloid precursor protein: retrograde and transcytotic transport in cultured neurons. J. Cell Biol. 129, 431, 1995. 23. Zuk, P.A. and Elferink, L.A. Rab15 mediates an early endocytic event in Chinese hamster ovary cells. J. Biol. Chem. 274, 22303, 1999.

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3

Strategies for Crystallizing the N-Terminal Growth Factor Domain of Amyloid Precursor Protein William J. McKinstry, Susanne C. Feil, Denise Galatis, Roberto Cappai and Michael W. Parker

CONTENTS Abstract 3.1 Introduction 3.2 Overview of Approach 3.3 Bioinformatics Analysis of APP 3.4 Biological Roles of GFD 3.5 Previous Crystallization Studies 3.6 Expression of Recombinant GFD in Pichia pastoris. 3.6.1 Materials 3.6.2 Method for Cloning 3.6.3 Method for Expression 3.7 Purification of GFD 3.7.1 Method 3.8 Crystallization of GFD 3.8.1 Materials 3.8.2 Method 3.9 Discussion Acknowledgments References

ABSTRACT The normal physiological roles of amyloid precursor protein (APP) remain largely unknown despite much research. A knowledge of APP function will not only provide insights into the genesis of Alzheimer’s disease, but may also prove vital in the development of an effective therapy. Here we describe our strategies for determining the three-dimensional atomic structure of APP, highlighting our work on the N-terminal growth factor domain. 0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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3.1 INTRODUCTION The normal physiological function of amyloid precursor protein (APP) remains largely unknown although a number of studies suggest that it acts as a cell surface receptor. APP shares a similar architecture,1 cellular orientation, and localization2,3 to known (type I) cell surface receptors. APP mutations associated with familial Alzheimer’s disease (FAD) cause constitutive activation of Go, a member of the heteromeric G protein family whose members serve as signal tranducers of cell surface receptors.4 It has been suggested that the FAD mutations may interfere with the possible dimerization of APP that leads to signal transduction, as may be the case with other cell surface receptors. The APP cytoplasmic domain binds to a number of proteins consistent with the possibility that APP functions as a receptor involved in signal transduction. For example, this domain binds to Fe65 protein, a protein related to oncogenic signal transducers.5 Other binding partners have been discovered including APP-BP1,6 X11,7 UV-DDB,8 Tip60,9 Numb,10 and ShcA/Grb2.11 Another series of studies have shown that an antibody directed toward the APP N-terminal domain stimulates G protein and MAP kinase activity.12,13 The antibody is presumably mimicking the action of a still-to-be-identified physiological ligand. The APP gene has been knocked out in mice, resulting in reduced body mass, reduced locomotor activity, and in some cases gliosis, indicating impaired neuronal function.14 In summary, current knowledge suggests APP is a potential Go-coupled receptor with ligand-regulated function, although the physiological roles of APP remain to be established. The uncertainty about the normal physiological roles of APP led us to embark on a structural investigation of the molecule. The availability of an atomic structure of APP might greatly aid studies directed toward understanding the normal functions of APP and might also prove useful for the design of novel therapeutics to combat Alzheimer’s disease.

3.2 OVERVIEW OF APPROACH APP represents a difficult target for crystallization. It is a heterogeneous membrane protein with multiple glycosylation, phosphorylation, and sulfation sites. Membrane proteins are very difficult to crystallize and only a small fraction of all protein crystal structures are of this type. In the case of APP, this problem can be circumvented by expressing APP fragments missing the transmembrane anchor. Highly heterogeneous proteins are difficult to crystallize and thus it is a common practice to minimize heterogeneity wherever possible. The heterogeneity of the protein bought about by glycosylation can be minimized or eliminated by glycosylases or mutating out potential glycosylation sites. Excessive phosphorylation can be overcome with the judicious use of phosphatases. APP is likely to be a highly mobile protein since it consists of numerous domains (see below). Multidomain proteins can be difficult to crystallize as protein flexibility can interfere with the crystallization process. A common strategy to overcome this problem is to target smaller fragments that might be more amenable to crystallization.

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The structure of the entire molecule can then be constructed by assembling the individual structures together.

3.3 BIOINFORMATICS ANALYSIS OF APP Before we embarked on crystallization studies, an in-depth analysis of the APP primary structure was carried out. Such an analysis can be very useful in deciding what fragments should be expressed and purified for crystallization trials. APP is a 90- to 130-kDa protein that is conserved among animal species and is expressed and secreted from a variety of tissues (see References 15 and 16 for reviews). There are at least 10 isoforms of APP due to alternative splicing of a single gene. The predominant isoform in neuronal tissues is a 695-amino acid protein. Its amino acid sequence reveals it is an integral membrane protein with a single transmembrane domain near the C-terminal end of the molecule (Figure 3.1). APP molecules from a variety of organisms have been sequenced. The locations of putative domains in the primary structure of APP have been determined based on extensive sequence alignments, secondary structure predictions, and database searches for similar sequences (Figure 3.1). The most highly conserved region of the molecule occurs at the N-terminal end, a cysteine-rich region that includes a heparin-binding domain (D1 or growth factor domain, GFD) and a metal-binding domain (D2 or copper binding domain, CuBD). All APP isoforms contain highly acidic domains; 45% of their residues are either Asp or Glu (D3). Two larger isoforms, APP751 and APP770, include an additional exon that encodes a domain with sequence similarity to a Kunitz protease inhibitor domain (KPI).17,18 The APP770 isoform also possesses a domain (D5) with similarity to the MRC OX-2 antigen, a neuronal membrane glycoprotein belonging to the immunoglobulin superfamily.18 Next is a 275-amino acid stretch that secondary structure predictions suggest consists of two domains: a highly helical domain (D6a) and a domain of little regular structure D1 growth factor

NH2-

D2 D3 CuBD acidic

115

100

D4 D5 KPI OX-2

56 19

D6 heparin binding

275

Αβ

D8 TM cytoplasmic

24 46 -COOH

β_ γ CHO membrane anchor clathrin secretase binding sites Go binding

FIGURE 3.1 Domain structure of APP. Domains are labeled D1 to D8 and the number of residues in each domain is indicated. The three-dimensional structure of GFD is shown as an alpha-carbon trace at the N-terminal end of the molecule. Known locations of carbohydrate attachment are denoted by “CHO.” Sites of proteolytic degradation are marked by Greek letters. The transmembrane (TM) is highlighted by dark shading. The location of the proteolytic breakdown product, Aβ, is also indicated.

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(D6b). The transmembrane (TM) region and cytoplasmic tail (D8) are located at the C-terminal end of the molecule.

3.4 BIOLOGICAL ROLES OF GFD The N-terminal region of APP, including GFD, has previously been shown to stimulate neurite outgrowth19 and regulate synaptogenesis.20 Furthermore, an antibody directed toward GFD stimulates G protein and MAP kinase activity.12,13 GFD has a known heparin-binding site.19 Heparan sulfate proteoglycans may play a key role in Alzheimer’s disease pathogenesis: exogenous heparin induces APP production and amyloidogenic secretion21 and a number of heparin-binding growth factors are expressed at increased levels in the brains of affected patients.22 Small heparin fragments have been proposed as possible drugs in preventing or retarding the disease.23 To learn more about this critical domain, we determined its three-dimensional structure by x-ray crystallography.

3.5 PREVIOUS CRYSTALLIZATION STUDIES We presented our original crystallization of GFD in 1999.24 We expressed a fragment of APP consisting of residues 18 to 350 and hence encompassing the putative N terminal domain, copper-binding domain and acidic-rich region of the molecule (Figure 3.1). Our intention was to crystallize the intact fragment but trace proteases generated a smaller fragment in the crystallization trials. Nevertheless, the smaller fragment yielded well diffracting crystals that led to the structural determination of GFD. N-terminal sequencing and mass spectrometry revealed the crystals consisted of a proteolytic breakdown product, residues 23 to 128. The final atomic model consisted of residues 28 to 123 indicating that residues 23 to 27 and 124 to 128 were too mobile to be seen in the electron density maps calculated from x-ray diffraction patterns generated from the crystals. To obtain better crystals of GFD, we decided to express GFD alone. We chose domain boundaries based on the crystal structure and omitted the most flexible regions to enhance the chances of obtaining high quality crystals (see below). We have now shown that expression of a fragment consisting of residues 28 to 123 crystallizes identically to the longer length fragment and the details are presented below.

3.6 EXPRESSION OF RECOMBINANT GFD IN PICHIA PASTORIS Recombinant secreted GFD (APP residues 28 to 123) was produced in the methylotrophic yeast Pichia pastoris using standard molecular biology protocols and P. pastoris protocols (Invitrogen, Carlsbad, CA). We chose the P. pastoris system as it offered high-level expression in a eukaryotic cell. In the first step, GFD was cloned into the P. pastoris expression plasmid pIC9 and then introduced into the P. pastoris cells as described below.

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3.6.1 MATERIALS The following media were used: 1. Yeast extract peptone dextrose (YPD) medium consisting of 1% (w/v) bacto yeast extract, 2% (w/v) peptone, and 2% (w/v) glucose. 2. Minimal methanol (MM) medium consisting of 1.34% (yeast nitrogen base without amino acids, 4 × 10-5% w/v) biotin and 2.0% (v/v) methanol. 3. Buffered methanol complex (BMMY) consisting of MM plus 1% (w/v) bacto yeast extract, 2% w/v peptone in 0.1 M phosphate buffer, pH 6.0. 4. Minimal dextrose (MD) plates, consisting of 1.34% (yeast nitrogen base without amino acids, 4 × 10-5% w/v) biotin, 1% dextrose, and 15 g agar for 1 liter. 5. Yeast extract peptone methanol (YPM) consisting of 1% (w/v) bacto yeast extract, 2% (w/v) peptone, and 3% (v/v) methanol.

3.6.2 METHOD

FOR

CLONING

1. The DNA encoding GFD is generated by polymerase chain reactin (PCR) using primers GGTCGACAAAAGAGAGGCTCTGCTGGCTGAACCCCAGATTG and GAATTCTTATACAAACTCACCAACTAAG. 2. The PCR product is cloned as a Xho1-EcoR1 fragment into the P. pastoris vector pIC9 (Invitrogen). 3. The construct is linearized with BglII prior to transformation into P. pastoris strain GS115 by electroporation. 4. A 5-ml quantity of GS115 is grown overnight in YPD in a 50-ml conical flask at 30oC. 5. On the following day, inoculate the overnight culture into 500 ml YPD in a 2-liter flask. Grow to OD600 of 1.3 to 1.5. 6. Centrifuge (1500 × g for 5 min) and resuspend cells in 500 ml ice cold water. 7. Centrifuge and resuspend in 250 ml ice-cold water. 8. Centrifuge and resuspend in 20 ml ice-cold 1 M sorbitol. 9. Centrifuge and resuspend in 1 ml ice-cold 1 M sorbitol. 10. Mix 80 µl of cells with 10 µg linearized pIC9-GFD DNA in a 0.2-cm electroporation cuvette on ice. Electroporate according to manufacturer’s recommended conditions for yeast. 11. Cells are spread onto MD plates and grown at 30oC for 2 to 5 days. 12. Colonies are grown in 5 ml YPD for 2 days, centrifuged and then grown in 1 ml BMMY for 2 days. 13. Expressing clones are identified by silver stain sodium dodecyl sulfate– polyacrylamide gel electrophoresis (SDS-PAGE) analysis of the culture supernatants. The expressing clones are then grown in culture to produce large amounts of GFD for crystallography and biological assays as described below.

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3.6.3

METHOD

FOR

EXPRESSION

1. Shaker flask cultures are grown by inoculating a suitable high-level expression clone into 500 ml YPD in a 2-liter baffled flask. 2. Cultures are grown for 48 hr at 3oC on an orbital shaker (250 rpm) to a cell density between 45 and 60 × 107 cells/ml (optical density of 15 to 20 at 600 nm). 3. Cells are harvested by centrifugation (2000 × g for 5 min), resuspended in 500 ml YPM, and grown for 48 hr at 30ºC on an orbital shaker (250 rpm). 4. Condition media containing the expressed GFD are harvested by centrifugation (14000 × g for 30 min) and filtered through a 0.45-µm filter.

3.7 PURIFICATION OF GFD GFD is known to bind heparin,19 and this property was used to purify the protein from the cell supernatants. Purification was performed using a Beckman 510 Protein Purification Workstation (Beckman Instruments, Fullerton, CA) fitted with a singlechannel wavelength detector tuned to 280 nm and a 1-cm path length analytical flow cell.

3.7.1 METHOD 1. A heparin–hyperD column (1.6 × 12 cm, Biosepra S.A., Cergy Saint Christophe, France) is equilibrated in 10 mM sodium phosphate buffer, pH 7.0, at a flow rate of 2.5 ml/min. The column is washed with equilibration buffer until the baseline returns to zero. 2. The supernatant is loaded directly onto the heparin–hyperD column. 3. Bound proteins are eluted with a 250-ml linear 0 to 2.0 M NaCl gradient in column equilibration buffer. 4. Next, 5-ml fractions are collected and analyzed by both SDS-PAGE and immunoblotting using a monoclonal antibody that recognizes this domain.12,13,25 5. Fractions containing GFD are pooled and buffer-exchanged into 20 mM Tris HCl, pH 8.0. 6. A QHyperD anion exchange column (4.6 × 100 mm, Biosepra S.A.) is equilibrated with 20 mM Tris HCl, pH 8.0. 7. The pooled fractions from the heparin column are loaded onto the QHyperD column and bound proteins eluted with a 50-ml linear gradient of NaCl (0 to 500 mM) in column equilibration buffer. 8. GFD elutes at a concentration of 50 mM NaCl. 9. The purified GFD will be >99% pure as judged by Coomassie blue staining of an overloaded SDS–polyacrylamide gel (Figure 3.2). 10. The purified GFD is concentrated between 4 and 5 mg/ml for crystallization trials. Care must be taken to maintain the purified GFD at 4oC because the protein readily forms microcrystals if allowed to warm up.

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FIGURE 3.2 GFD purity as assessed by SDS-PAGE and Coomassie blue staining. Molecular weight markers are shown in the left column, with their masses in kDa.

3.8 CRYSTALLIZATION OF GFD Proteins can be made to crystallize by the addition of certain precipitants such as salts and organic solvents, most commonly ammonium sulfate or polyethylene glycol, under unusually precise conditions of pH, temperature and protein concentration. Many factors can influence successful crystallization including protein and precipitant concentrations, ionic strength, vibration, protein flexibility, protein purity, small molecule additives, temperature, and so on. The detailed physics behind crystallization are not well understood. The process is usually considered in terms of phase diagrams where the vertical axis corresponds to the protein solubility and the horizontal axis refers to some experimental parameter such as pH or precipitant concentration. Consider the behavior of a typical protein solution. At low protein and precipitant concentrations, the protein stays in solution (i.e., it is undersaturated). As the concentration of protein or precipitant increases, the protein becomes less soluble until supersaturation occurs whereby the protein comes out of solution as either an amorphous precipitate or as ordered crystals. All crystallization experiments for APP were conducted using the hanging drop vapor diffusion method.26 The wells of a tissue culture tray were filled with a precipitant solution and a mixture of protein and precipitant solution was applied to the surface of a coverslip that was then placed over the well of a culture dish, the drop facing down (Figure 3.3). The protein and precipitant in the drop slowly become more concentrated and a point is reached when the protein reaches supersaturation and will form an amorphous precipitate or crystal nuclei. When the precipitate or crystals form, the protein concentration decreases. Crystals may begin to form at any time, from the start of the equilibration process until long after equilibrium has been reached, and may form after precipitation of the protein has occurred.

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Hanging Drop

Cover Slip

Reservoir

FIGURE 3.3 Crystallization by the vapor diffusion hanging drop method. (Courtesy of Geoffrey Kong.)

The point of supersaturation is governed by the protein and the type of precipitant used. The rate of equilibration is governed by temperature, drop size, and the type of precipitant used. The protein in the drop only becomes more concentrated if the precipitant or salt concentration is higher in the well. It is not possible to determine a priori what conditions will be required for the crystallization of a new protein. Many possible crystallization conditions could have been tested and trials were carried out using large screens under many different conditions. The crystallization protocol for GFD is explained below.

3.8.1 MATERIALS 1, Tissue culture plates are available from ICN Biochemicals, Inc. (Aurora, OH). 2. Chemicals for crystallization may be obtained from Fluka (Buchs, Switzerland) or Sigma-Aldrich (Sydney, Australia). 3. Amicon protein microconcentrators may be obtained from Amicon, Inc. (Beverly, MA).

3.8.2 METHOD 1. The purified protein is dialyzed into 5 mM Tris HCl buffer, pH 7.5. 2. The protein is concentrated to 10 mg/ml in Amicon concentrators. 3. Take a tissue culture plate and fill the reservoirs with 1 ml of solutions containing from 18 to 26% (w/w) PEG 10K (steps of 2%) and 100 mM HEPES buffer, ranging from pH 7.0 to 8.0 (steps of 0.5 pH units). 4. Grease the rims of the wells with petroleum jelly. 5. Mix 2 µl of protein with 2 µl of reservoir solution on a cover slip and hang the cover slip over 1 ml of reservoir solution. 6. Store the trays in a constant temperature room set to 22°C. 7. Crystals should appear in a number of the drops after 4 days. 8. The crystals take 2 weeks to grow to maximal dimensions of approximately 0.2 × 0.2 × 0.4 mm (Figure 3.4).

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FIGURE 3.4 Crystals of GFD. The largest crystal is 0.3 mm in its longest dimension.

3.9 DISCUSSION The availability of well-diffracting crystals was the vital first step in determining the three-dimensional atomic structure of GFD by x-ray crystallography.26 GFD was found to adopt a compact, globular fold consisting of nine β-strands and one α-helix tethered together by three disulfide bridges (Cys 38 to Cys 62; Cys 73 to Cys 117; Cys 98 to Cys 105; see Figure 3.5). Sequence alignments of APP orthologues and paralogues show their GFD regions are well conserved with sequence identities ranging from 36 to 84%; all the cysteine residues are strictly conserved. This suggests that the fold described here is maintained across the APP family. An electrostatic calculation based on the three-dimensional structure demonstrated a highly positively charged surface on one side of the domain including a peptide region, residues 96 to 110, that was previously identified as part of a heparinbinding site.19 Maintenance of the disulfide bridge in this region is critical for neurite outgrowth19 and activation of MAP kinase,27 suggesting that the conformation of the loop is important. The surface is dominated by the β-hairpin loop (residues 96 to 110) representing the most mobile region of the structure. The overall fold of the GFD did not resemble any protein of known three-dimensional structure. However, like APP, a number of growth factors also possess disulfide-bonded β-hairpin loops implicated in proteoglycan binding. These include midkine,28 hepatocyte growth factor29 and vascular endothelial growth factor.30 In all cases, the loop is long, flexible, and highly charged with basic residues. These properties appear ideal for binding heparin oligosaccharides where the flexibility would allow induced fit binding via the positively charged residues around the sulfate moieties of the carbohydrate.

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FIGURE 3.5 Structure of GFD. A ribbon diagram indicating the location of secondary structure with helices as coils and β-strands as arrows. The disulfide bridges are shown in ball-and-stick form and the tip of the β-hairpin loop is indicated. This figure was drawn with MOLSCRIPT.34

The extracellular domain of APP has been shown to be a potent mediator of thyrocyte proliferation31 and can potentiate the phosphorylation of the tyrosine kinase (trkA) receptor caused by nerve growth factor binding.32 The N-terminal cysteinerich region can inhibit platelet activation33 emphasizing the role of this region in modulating cellular pathways and function. Presumably, the growth-promoting activity of APP is expressed after it is released from membranes through the action of secretases. In conclusion, the structural similarities to some growth factors in concert with the known growth promoting properties of APP and its N-terminal domain led to the conclusion that GFD can be classified as a new member of the cysteine-rich growth factor superfamily.

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ACKNOWLEDGMENTS This work was supported by grants from the National Health and Medical Research Council of Australia to R.C. and M.W.P. W.J.M. is a NHMRC Industry Fellow and M.W.P. is a NHMRC Senior Principal Research Fellow.

REFERENCES 1. Kang, J. et al. The precursor of Alzheimer’s disease amyloid A4 protein resembles a cell-surface receptor. Nature 325, 733, 1987. 2. Weidemann, A. et al. Identification, biogenesis, and localization of precursors of Alzheimer’s disease A4 amyloid protein. Cell 57, 115, 1989. 3. Schubert, W. et al. Localization of Alzheimer beta A4 amyloid precursor protein at central and peripheral synaptic sites. Brain Res. 563, 184, 1991. 4. Okamoto, T. et al. Intrinsic signaling function of APP as a novel target of three V642 mutations linked to familial Alzheimer’s disease. EMBO J. 15, 3769, 1996. 5. Zambrano, N. et al. Interaction of the phosphotyrosine interaction/phosphotyrosine binding-related domains of Fe65 with wild-type and mutant Alzheimer’s beta-amyloid precursor proteins. J. Biol. Chem. 272, 6399, 1997. 6. Chow, N. et al. APP-BP1, a novel protein that binds to the carboxyl-terminal region of the amyloid precursor protein. J. Biol. Chem. 271, 11339, 1996. 7. Borg, J.P. et al. The phosphotyrosine interaction domains of X11 and FE65 bind to distinct sites on the YENPTY motif of amyloid precursor protein. Mol. Cell. Biol. 16, 6229, 1996. 8. Watanabe, T. et al. A 127-kDa protein (UV-DDB) binds to the cytoplasmic domain of the Alzheimer’s amyloid precursor protein. J. Neurochem. 72, 549, 1999. 9. Cao, X. and Südhof., T.C. A transcriptionally active complex of APP with Fe65 and histone acetyltransferase Tip60. Science 293, 115, 2001. 10. Roncarati, R. et al. The gamma-secretase-generated intracellular domain of betaamyloid precursor protein binds Numb and inhibits Notch signaling. Proc. Natl. Acad. Sci. USA 99, 7102, 2002. 11. Russo, C. et al. Signal transduction through tyrosine-phosphorylated C-terminal fragments of amyloid precursor protein via an enhanced interaction with Shc/Grb2 adaptor proteins in reactive astrocytes of Alzheimer’s disease brain. J. Biol. Chem. 277, 35282, 2002. 12. Okamoto, T. et al. Ligand-dependent G protein coupling function of amyloid transmembrane precursor. J. Biol. Chem. 270, 4205, 1995. 13. Murayama, Y. et al. Cell surface receptor function of amyloid precursor protein that activates Ser/Thr kinases. Gerontology 42, 2, 1996. 14. Zheng, H. et al. beta-Amyloid precursor protein-deficient mice show reactive gliosis and decreased locomotor activity. Cell 81, 525, 1995. 15. Hendriks, L. and Van Broeckhoven, C. A beta A4 amyloid precursor protein gene and Alzheimer’s disease. Eur. J. Biochem. 237, 6, 1996. 16. Mattson, M.P. Cellular actions of beta-amyloid precursor protein and its soluble and fibrillogenic derivatives. Physiol. Revs. 77, 1081, 1997. 17. Kitaguchi, N. et al. Novel precursor of Alzheimer’s disease amyloid protein shows protease inhibitory activity. Nature 331, 530, 1988. 18. Ponte, P. et al. A new A4 amyloid mRNA contains a domain homologous to serine proteinase inhibitors. Nature 331, 525, 1988.

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19. Small, D.H et al. A heparin-binding domain in the amyloid protein precursor of Alzheimer’s disease is involved in the regulation of neurite outgrowth. J. Neurosci. 14, 2117, 1994. 20. Morimoto, T. et al. Involvement of amyloid precursor protein in functional synapse formation in cultured hippocampal neurons. J. Neurosci. Res. 51, 185, 1998. 21, Leveugle, B. et al. Heparin promotes beta-secretase cleavage of the Alzheimer’s amyloid precursor protein. Neurochem. Int. 30, 543, 1997. 22. Fenton, H. et al. Hepatocyte growth factor (HGF/SF) in Alzheimer’s disease. Brain Res. 779, 262, 1998. 23. Leveugle, B. et al. Heparin oligosaccharides that pass the blood–brain barrier inhibit beta-amyloid precursor protein secretion and heparin binding to beta-amyloid peptide. J. Neurochem. 70, 736, 1998. 24. Rossjohn, J. et al. Crystal structure of the N-terminal, growth factor-like domain of Alzheimer amyloid precursor protein. Nature Struct. Biol. 6, 327, 1999. 25. Hilbich, C. et al. Amyloid-like properties of peptides flanking the epitope of amyloid precursor protein-specific monoclonal antibody 22C11. J. Biol. Chem. 268, 26571, 1993. 26. McPherson, A., Crystallization of Biological Macromolecules, Cold Spring Harbor Laboratory Press, New York, 1999. 27. Greenberg, S.M. et al. Amino-terminal region of the beta-amyloid precursor protein activates mitogen-activated protein kinase. Neurosci. Lett. 198, 52, 1995. 28. Iwasaki, W. et al. Solution structure of midkine, a new heparin-binding growth factor. EMBO J. 16, 6936, 1997. 29. Zhou, H. et al. The solution structure of the N-terminal domain of hepatocyte growth factor reveals a potential heparin-binding site. Structure 6, 109, 1998. 30. Fairbrother, W.J. et al. Solution structure of the heparin-binding domain of vascular endothelial growth factor. Structure 6, 637, 1998. 31. Pietrzik, C.U. et al. From differentiation to proliferation: the secretory amyloid precursor protein as a local mediator of growth in thyroid epithelial cells. Proc. Natl. Acad. Sci. USA 95, 1770, 1998. 32. Akar, C.A. and Wallace, W.C. Amyloid precursor protein modulates the interaction of nerve growth factor with p75 receptor and potentiates its activation of trkA phosphorylation. Mol. Brain. Res. 56, 125, 1998. 33. Henry A. et al. Inhibition of platelet activation by the Alzheimer’s disease amyloid precursor protein. Br. J. Haematol. 103, 402, 1998. 34. Kraulis, P.J. MOLSCRIPT: a program to produce both detailed and schematic plots of proteins. J. Appl. Crystallogr. 24, 946, 1991.

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4

Analysis of Amyloid Precursor Protein Processing Protease β-Secretase: Tools for Memapsin 2 (β-Secretase) Inhibition Studies Gerald Koelsch, Vajira Weerasena, Dongwoo Shin, Arun K. Ghosh, and Jordan Tang

CONTENTS 4.1 4.2 4.3

Introduction Assay of Memapsin 2 Activity Synthesis of Memapsin 2 Inhibitors Based on APP Sequence 4.3.1 N-(tert-butoxycarbonyl)-L-leucine-N′-methoxy-N′-methylamide (3) 4.3.2 N-(tert-butoxycarbonyl)-L-leucinal (4) 4.3.3 Ethyl (4S,5S)- and (4R,5S)-5-[(tert-butoxycarbonyl)amino]-4hydroxy-7-methyloct-2-ynoate (5) 4.3.4 (5S,1′S)-5-[1′-[(tert-Butoxycarbonyl)amino]-3′-methylbutyl]dihydrofuran-2(3H)-one (7) 4.3.5 (3R,5S,1′S)-5-[1′-[(tert-butoxycarbonyl)amino)]-3′-methylbutyl]-3 methyl dihydrofuran-2(3H)-one (8) 4.3.6 (2R,4S,5S)-5-[(tert-Butoxycarbonyl)amino]-4-[(tertbutyldimethylsilyl)oxy ]-2,7-dimethyloctanoicacid (9) 4.3.7 (2R,4S,5S)-5-[(fluorenylmethyloxycarbonyl)amino]-4-[(tertbutyldimethyl silyl)oxy]-2,7-dimethyloctanoic acid (10) 4.3.8 Coupling of Di-Isostere in Solid-Phase Peptide Synthesis 4.4 Determination of Inhibition Constants 4.5 Summary References

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4.1 INTRODUCTION

Fluorescence Intensity

Memapsin 2 (BACE, ASP-2) is the membrane-anchored aspartic protease that processes APP at the β-secretase site.1–5 The resulting C terminal fragment, C99, is further processed by γ-secretase to produce amyloid-β (Aβ) peptide.6 Since Aβ is intimately related to the pathogenesis of Alzheimer’s disease, a great deal of interest surrounds the design and testing of memapsin 2 inhibitors. The first potent transition state inhibitor of memapsin 2, OM99-2 (Figure 4.1), was designed based on a slightly modified sequence around the β-secretase processing site of APP Swedish mutant.7 This inhibitor had a Ki of 1.6 nM. When the complete subsite specificity of memapsin 2 was determined,8 the most preferred residues were used to design OM00-3 (Figure 4.2) which had a Ki of 0.3 nM.9 The crystal structures of these inhibitors bound to the catalytic unit of memapsin 2 have been reported.10,11 These structures define the interactions of the protease active sites with the inhibitors and also predict the locations of the APP substrate binding positions during the hydrolysis. Memapsin 2 is a type I transmembrane protein with the catalytic domain located on the lumenal face of the plasma membrane.1–5 Likewise, its most notorious substrate, APP, has the same topology as the β-secretase cleavage site located within 28 amino acids of the plasma membrane surface. In vivo memapsin 2 may be regulated by factors including expression, post-translational modifications and membrane component compositions such as “lipid rafts” and endocytotic and vesicular trafficking to acidic compartments such as endosomes. Nonetheless, small peptide substrates representing the β-secretase cleavage site in APP are capably cleaved by memapsin 2,5,12 demonstrating the independence of in vivo biochemical regulation from its fundamental proteolytic function. Similarly, retroviral proteases demonstrate complex in vivo characteristics, requiring dimerization in the form of gag-pol precursor proteins with subsequent activation from these precursor multiproteins. These proteases are still able to cleave peptide substrates in vitro and autoprocess a mini-precursor form of the protease.13,14 Despite the complexity of the in vivo environment of the retroviral proteases, clearly 0.35

0.30

0.25

0.20 0

50

100

150

200

Time (sec)

FIGURE 4.1 Continuous fluorescence assay of memapsin 2 activity using an Mca/Dnp internally quenched fluorogenic substrate. Memapsin 2 was incubated with substrate (3 µM) at pH 4.0, 37˚C and emission at 393 nm was monitored continuously with excitation at 328 nm. Increased fluorescence intensity over time indicates proteolysis of the substrate. Linear regression is used to determine the initial velocity (V0) of the uninhibited enzyme.

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the development of potent HIV inhibitors has been successful using peptide-based in vitro enzyme assays. Thus it is likely that memapsin 2 inhibitors with pharmaceutical application for modulation of Aβ production may be developed from the assay of small peptide substrates that model the fundamental proteolytic function, superseding the complexity of the memapsin 2 function in vivo. The assay of memapsin 2 activity, syntheses of transition-state memapsin 2 inhibitors, and measurements of their potencies are described in the following sections.

4.2 ASSAY OF MEMAPSIN 2 ACTIVITY Continuous assay of memapsin 2 activity is a necessary prelude to the measure of its inhibition and permits rapid analysis of inhibition potency and determination of inhibition constants. An internally quenched fluorogenic substrate for memapsin 2 was designed to mimic the β-secretase cleavage site of APP (substrate FS-2 in Reference 12) with the sequence Mca–Ser–Glu–Val–Asn–Leu–Asp–Ala–Glu–Phe– Lys(Dnp)–NH2 [Mca–(Asn670, Leu671)–APP770 (667–675)–Lys(Dnp) amide] and is available from Bachem (Torrance, CA, #2485). It contains the (7-methoxycoumarin4-yl)acetyl fluorophore (Mca) at the amino terminus (effectively at the P6 position) and the quenching chromophore group N-2,4-dinitrophenyl (Dnp) attached to the ε-amino group of Lys in the P′5 position. Upon excitation of the Mca group at 328 nm, energy is transferred to the Dnp group with limited photon emission detectible at 393 nm (fluorescence resonance energy transfer or FRET). Cleavage of the intervening peptide results in diffusion of the two products, each containing a respective fluorophore and quenching chromophore group. This permits the excitation of the Mca group of the N-terminal product, resulting in unquenched emission at a wavelength of 393 nm (λex = 328 nm, λem = 393 nm). Increased fluorescence intensity at this excitation–emission wavelength pair allows continuous monitoring of proteolytic activity (Figure 4.1). Cleavage at the β-secretase site alone by memapsin 2 was confirmed by mass spectrometry.12 The assay of β-secretase activity using the Mca–Dnp substrate is accomplished by the addition of 1.75 ml 0.1 M sodium acetate, pH 4.0, to an aliquot of dimethyl sulfoxide (DMSO) in a 1.0 × 1.0-cm quartz cuvette thermostatted cell holder preequilibrated to 37˚C. The DMSO is added to the aliquot of sodium acetate such that the final DMSO concentration is 10% (including the amounts of substrate and inhibitor to be added, which are dissolved in DMSO). Memapsin 2 activity versus substrate FS-2 was found to be optimal at this concentration of DMSO.12 Memapsin 25,12 enzyme stock (typically 6 µM, 50 µl aliquot) is added, followed by substrate (20 µl of 300 µM stock FS-2 in DMSO; see Reference 12) to initiate the reaction (2 ml total volume). Fluorescence intensity over a 5-min period was monitored with excitation at 328 nm and emission at 393 nm, using a detector voltage of 700 V on an Aminco Bowman luminescence spectrometer. Fit of the linear portion of the signal to a linear model produces a typical signal of 10-3 fluorescence units per second (FU/sec). A typical time trace for hydrolysis of FS-2 by memapsin 2 is shown in Figure 4.1. The determined rate of reaction is proportional to the rate of cleavage of molar amounts of substrate per unit volume, but requires conversion of the observed initial

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velocity (in FU/sec) using the response factor of known concentrations of fluorophore and the ratio of free flourophore to that of the cleavage product, due to the inner filter effect of intermolecular quenching.15 However, this correction is not necessary in the determination of the inhibition constant Kiapp (see below) because relative initial velocities are determined at a fixed substrate concentration.

4.3 SYNTHESIS OF MEMAPSIN 2 INHIBITORS BASED ON APP SEQUENCE Transition state theory indicates that an enzyme will bind most tightly to its substrate when the substrate adopts a conformation approximating the transition state in the progression toward product formation. Thus the enzyme induces or stabilizes the substrate in that conformation, lowering the activation energy to permit more frequent progression to product. Inhibition of this process is therefore best accomplished with compounds that mimic the transition state of the scissile bond and are thus potent inhibitors of β-secretase.16 Peptide-based inhibitors of β-secretase have been described by our laboratory.8,9,17 The synthesis of OM99-2 and OM00-3 consists of two major steps. The first is the synthesis of a Leu*Ala dipeptide transition state isostere (that will be referred to as the di-isostere; the asterisk represents the hydroxyethylene moiety) with appropriate protective groups. The second step is to use solid-phase peptide synthesis to incorporate the di-isostere into the inhibitor. The synthesis of the Leu*Ala di-isostere using commercial BOC-leucine as a starting material is outlined in Figure 4.2. The Fmoc-protected di-isostere (compound 10 in Figure 4.2) is inserted in a coupling step as for other amino acid residues in the peptide synthesis. In this manner, it is possible to develop inhibitors to exploit the substrate specificity of memapsin 29 by incorporating various standard or nonstandard amino acids into the peptide. Thus incorporation of the di-isostere in solidphase peptide synthesis creates a molecule endowed with the potential to inhibit memapsin 2 activity. Detailed steps for the synthesis of the Leu*Ala di-isostere are discussed in the following sections. The compound numbers appearing in parentheses in the headings below correspond to Figure 4.2.

4.3.1 N-(TERT-BUTOXYCARBONYL)-L-LEUCINE-N′-METHOXY-N′-METHYLAMIDE (3) To a stirred solution of N,O-dimethylhydroxyamine hydrochloride (5.52 g, 56.6 mmol) in dry dichloromethane (25 mL) under N2 atmosphere at 0˚C, 1-methylpiperidine (6.9 mL, 56.6 mmol) is added dropwise. The resulting mixture is stirred at 0˚C for 30 min. In a separate flask, N-(tert-butyloxycarbonyl)-L-leucine (BOC-leucine, 2) (11.9 g, 51.4 mmol) is dissolved in a mixture of THF (tetrahydrofuran, 45 mL) and dichloromethane (180 mL) under N2 atmosphere. The resulting solution is cooled to –20˚C. To this solution is added 1-methylpiperidine (6.9 mL, 56.6 mmol) followed by isobutyl chloroformate (7.3 mL, 56.6 mmol). The resulting mixture is stirred for 5 min at –20˚C and the above solution of N,O-dimethylhydroxyamine is added to it. The reaction mixture is kept at –20oC for

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FIGURE 4.2 Scheme for synthesis of Leu*Ala di-isostere. Letters accompanying arrows refer to reagents and conditions: (a) LiAlH4, Et2O, –40oC, 30 min (86%); (b) LDA, HC≡C-CO2Et, THF, –78oC, 30 min, then 4, –78oC, 1 hr (42%); (c) H2, Pd-BaSO4, EtOAc; (d) AcOH, PhMe, reflux, 6hr (74%); (e) LiHMDS, MeI, THF, –78oC, 20 min (76%); (f) aqueous LiOH, THFH2O, 23oC, 10 hr; (g) TBDMSCl, imidazole, DMF, 24 hr (90%); (h) CF3CO2H, CH2Cl2, 0oC, 1.5 hr; (i) Fmoc-OSu, aqueous NaHCO3, dioxane, 23oC, 8 hr (61%).

30 min and then warmed to 23˚C. The reaction is quenched with water and the layers are separated. The aqueous layer is extracted with dichloromethane (3 × 100 mL). The combined organic layers are washed with 10% citric acid, saturated sodium bicarbonate, and brine. The organic layer is dried over anhydrous Na2SO4 and concentrated under the reduced pressure. The residue is purified by flash silica gel chromatography (25% ethyl acetate–hexane) to yield the title compound 3 as a pale yellow oil.

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4.3.2 N-(TERT-BUTOXYCARBONYL)-L-LEUCINAL (4) To a stirred suspension of lithium aluminum hydride (770 mg, 20.3 mmol) in dry diethyl ether (60 mL) at –40oC under N2 atmosphere is added N-tert-butyloxycarbonyl-L-leucine-N′-methoxy-N′-methylamide (5.05 g, 18.4 mmol) in diethyl ether (20 mL). The resulting reaction mixture is stirred for 30 min, after which the reaction is quenched with 10% NaHSO4 solution (30 mL). The resulting reaction mixture is then warmed to 23˚C and stirred at that temperature for 30 min. The resulting solution is filtered and the filter cake is washed with two portions of diethyl ether. The combined organic layers are washed with saturated sodium bicarbonate and brine and dried over anhydrous MgSO4. Evaporation of the solvent under reduced pressure yields the title aldehyde 4 (3.41 g) as a pale yellow oil. The resulting aldehyde is used immediately without further purification.

4.3.3 ETHYL (4S,5S)- AND (4R,5S)-5-[(TERT-BUTOXYCARBONYL)AMINO]4-HYDROXY-7-METHYLOCT-2-YNOATE (5) To a stirred solution of diisopropylamine (1.1 mL, 7.9 mmol) in dry THF (60 mL) at 0oC under N2 atmosphere is added n-BuLi (1.6 M in hexane, 4.95 mL, 7.9 mmol) dropwise. The resulting solution is stirred at 0oC for 5 min and then warmed to 23˚C and stirred for 15 min. The mixture is cooled to –78oC and ethyl propiolate (801 µL) in THF (2 mL) is added dropwise over a period of 5 min. The mixture is stirred for 30 min, after which N-BOC-L-leucinal 4 (1.55 g, 7.2 mmol) in 8 mL of dry THF is added. The resulting mixture is stirred at –78oC for 1 hr, after which the reaction is quenched with acetic acid (5 mL) in THF (20 mL). The reaction mixture is warmed to 23˚C and brine solution is added. The layers are separated and the organic layer is washed with saturated sodium bicarbonate and dried over Na2SO4. Evaporation of the solvent under reduced pressure provides a residue that is purified by flash silica gel chromatography (15% ethyl acetate–hexane) to afford a 3:1 mixture of acetylenic alcohols 5.

4.3.4 (5S,1′′S)-5-[1′′-[(TERT-BUTOXYCARBONYL)AMINO]-3′′METHYLBUTYL]-DIHYDROFURAN-2(3H)-ONE (7) To a stirred solution of the above mixture of acetylenic alcohols (1.73 g, 5.5 mmol) in ethyl acetate (20 mL) is added 5% Pd–BaSO4 (1 g). The resulting mixture is hydrogenated at 50 psi for 1.5 hr. After this period, the catalyst is filtered off through a plug of Celite and the filtrate concentrated under reduced pressure. The residue is dissolved in toluene (20 mL) and acetic acid (100 µL). The reaction mixture is refluxed for 6 hr, after which the reaction is cooled to 23˚C and the solvent is evaporated to produce a residue purified by flash silica gel chromatography (40% diethyl ether–hexane) to yield the (5S,1S′)-γ-lactone 7 (0.94 g, 62.8%) and the (5R,1S′)-γlactone 6 (0.16 g, 10.7%).

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4.3.5 (3R,5S,1′′S)-5-[1′′-[(TERT-BUTOXYCARBONYL)AMINO)]3′′-METHYLBUTYL]-3-METHYL DIHYDROFURAN-2(3H)-ONE (8) To a stirred solution of the lactone 7 (451.8 mg, 1.67 mmol) in dry THF (8 mL) at –78oC under N2 atmosphere is added lithium hexamethyldisilazane (3.67 mL, 1.0 M in THF) over a period of 3 min. The resulting mixture is stirred at –78oC for 30 min to generate lithium enolate. After this period, CH3I (228 µL) is added dropwise and the resulting mixture stirred at –78˚C for 20 min. The reaction is quenched with saturated aqueous NH4Cl solution and allowed to warm to 23˚C. The reaction mixture is concentrated under reduced pressure and the residue extracted with ethyl acetate (3 × 100 mL). The combined organic layers are washed with brine and dried over anhydrous Na2SO4. Evaporation of the solvent affords a residue that is purified by silica gel chromatography (15% ethyl acetate–hexane) to furnish the alkylated lactone 8 (0.36 g, 76%) as an amorphous solid.

4.3.6 (2R,4S,5S)-5-[(TERT-BUTOXYCARBONYL)AMINO]-4-[(TERTBUTYLDIMETHYLSILYL)OXY ]-2,7-DIMETHYLOCTANOICACID (9) To a stirred solution of lactone 8 (0.33 g, 1.17 mmol) in THF (2 mL) is added 1 N aqueous LiOH solution (5.8 mL). The resulting mixture is stirred at 23˚C for 10 hr, after which the reaction mixture is concentrated under reduced pressure and the remaining aqueous residue cooled to 0oC and acidified with 25% citric acid solution to pH 4. The resulting acidic solution is extracted with ethyl acetate (3 × 50 mL). The combined organic layers are washed with brine, dried over Na2SO4 and concentrated to yield the corresponding hydroxy acid (330 mg) as a white foam. This hydroxy acid is used directly for the next reaction without further purification. To the hydroxy acid (330 mg, 1.1 mmol) in anhydrous DMF is added imidazole (1.59 g, 23.34 mmol) and tert-butyldimethylchlorosilane (1.76 g, 11.67 mmol). The resulting mixture is stirred at 23˚C for 24 hr. After this period, MeOH (4 mL) is added and the mixture stirred for 1 hr. The mixture is then diluted with 25% citric acid (20 mL) and extracted with ethyl acetate (3 × 20 mL). The combined extracts are washed with water and brine and dried over anhydrous Na2SO4. Evaporation of the solvent produces a viscous oil that is purified by flash chromatography over silica gel (35% ethyl acetate–hexane) to afford the silyl protected acid 9.

4.3.7 (2R,4S,5S)-5-[(FLUORENYLMETHYLOXYCARBONYL)AMINO]4-[(TERT-BUTYLDIMETHYL SILYL)OXY]-2,7-DIMETHYLOCTANOIC

ACID

(10)

To a stirred solution of the acid 9 (0.17 g, 0.41 mmol) in dichloromethane (2 mL) at 0oC is added trifluoroacetic acid (500 µL). The resulting mixture is stirred at 0oC for 1 hr and an additional 500 µL of trifluoroacetic acid is added to the reaction mixture. The mixture is stirred for an additional 30 min and the progress of the reaction monitored by thin layer chromatography (TLC). After this period, the solvents are carefully removed under reduced pressure at a bath temperature not

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exceeding 5oC. The residue is dissolved in dioxane (3 mL) and NaHCO3 (300 mg) in 5 mL of H2O. To this solution is added Fmoc-succinimide (166.5 mg, 0.49 mmol) in 5 mL of dioxane. The resulting mixture is stirred at 23˚C for 8 hr. The mixure is then diluted with H2O (5 mL) and acidified with 25% aqueous citric acid to pH 4. The acidic solution is extracted with ethyl acetate (3 × 50 mL). The combined extracts are washed with brine, dried over Na2SO4, and concentrated under reduced pressure to yield a viscous oil residue. Purification of the residue by flash chromatography over silica gel affords the Fmoc-protected acid 10.

4.3.8 COUPLING

OF

DI-ISOSTERE

IN

SOLID-PHASE PEPTIDE SYNTHESIS

The solid-phase peptide synthesis protocol may be utilized for the synthesis of inhibitors. The synthesis is initiated with Wang resin (0.3 mmol) precoupled with a carboxy terminal amino acid of choice and capped with a N-9-fluorenylmethyloxycarbonyl(Fmoc) alpha amino-protecting group. The coupling of N-Fmoc-Ala, isostere 10 is accomplished as follows. The removal of the N-Fmoc group from the peptide chain downstream of the di-isostere is carried out in 20% piperidine in dimethylforamide for 15 min. The peptide coupling reaction is accomplished with 2-(1Hbenzotriazol-1-yl) 1,1,3,3-tetramethyluronium tetrafluoroborate, 1-hydroxybenzotriazole and diisopropylethylamine (3.3 equivalents each) in N-methyl pyrolidine. After the di-isostere coupling, other Fmoc-protected derivatives are coupled. After the last residue coupling step, the peptide is cleaved from the solid-state resin using 95% trifluoroacetic acid, which also removes all the side chain-protecting groups including the silyl-group of 10. The inhibitors are purified in reversed-phase high performance liquid chromatography (HPLC) using a C18 column equilibrated in 0.1% trifluoroacetic acid in H2O with a linear gradient of acetonitrile from 0 to 25% over 25 min.

4.4 DETERMINATION OF INHIBITION CONSTANTS Because the inhibitors of interest are very potent and have Ki values in the nM range, the inhibition constants cannot be determined accurately by conventional steadystate kinetics. The inhibition constant Kiapp is determined from nonlinear regression of the model of Bieth.18 Proteolytic activity in the presence of inhibitor (Vi) and full activity free of inhibitor (V0) are measured with a constant concentration of the enzyme. The mixture of the enzyme and inhibitor is pre-equilibrated for 20 min and the reaction is initiated by the addition of the substrate. The relative activity (a) is the ratio of Vi /V0 and is determined at various concentrations of inhibitor. The apparent inhibition constant, Kiapp, may be determined from a plot of relative activity (a) versus [I] (Figure 4.3) based on Equation 1:

(

)

2

1 − I 0 +  E 0 + K iapp − I 0 +  E 0 + K iapp − 4 I 0  E 0 a= 2  E 0

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Relative Activity

1.2 1.0 0.8 0.6 0.4 0.2 0.0 0

200

400

[I], nM

FIGURE 4.3 Determination of Kiapp from a profile of relative activity vs. inhibitor concentration. Initial velocities (Vi) were determined at various concentrations of inhibitor and expressed relative to the initial velocity of uninhibited control reaction (V0) (solid symbols). Nonlinear regression of the data (solid line) with Equation 1 determines Kiapp.

Nonlinear regression of the data with the above equation to obtain Kiapp may be accomplished using GraFit.19 Typically 5 to 10 determinations of relative activity over a range of [I] will produce an error of the fit to ±5 to 10%. Sensitivity of the model to [E]0 requires that the concentration of the enzyme stock be determined accurately. Simple conversion of optical density measurements at 280 nm may not provide accurate information. Rather it is preferred to determine the number of active sites by titration using a tight-binding inhibitor.20 The data from determination of [E]0 by this method may also be used to confirm the Kiapp value for the titrating inhibitor, using additional determinations at [I] beyond the range of the linear relationship between a and [I]. The obtained Kiapp measurement may be dependent upon substrate concentration [12] and may be corrected by the relationship:

(

K iapp = K i 1 + S  K m

)

4.5 SUMMARY Inhibitors of aspartic proteases may be developed from peptide substrate templates. In the case of memapsin 2, by employing the basic sequence of a good substrate from the APP Swedish mutant and the principle of a transition-state mimic,21 potent inhibitors like OM99-2 can be developed. In this chapter we provide basic methods for such work. The synthesis of a transition-state isostere in the place of the scissile peptide bond is essential for a tight-binding transition-state inhibitor. The synthetic procedure of a blocked two-residue isostere has the advantage that it can be used in standard solid-phase peptide synthesis of a long inhibitor. This approach is suitable to explore diverse amino acid sequences, including nonstandard amino acids.8,17 The second essential aspect in inhibitor development is the kinetic assay for inhibition potency. A fluorogenic substrate based on a slightly modified sequence of the APP Swedish mutant is described. This assay,12 based on the principle of fluorescence resonance energy transfer, is very sensitive and has been reliably used to determine competitive inhibition constants using a model for tight-binding inhibitors.8,17

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REFERENCES 1. Vassar, R. et al. β-Secretase cleavage of Alzheimer’s amyloid precursor protein by the transmembrane aspartic protease BACE. Science 286, 735–741, 1999. 2. Sinha, S. et al. Purification and cloning of amyloid precursor protein β-secretase from human brain. Nature 402, 537–540, 1999. 3. Yan, R. et al. Membrane-anchored aspartyl protease with Alzheimer’s disease β-secretase activity. Nature 402, 533–537, 1999. 4. Hussain, I. et al. Identification of a novel aspartic protease (Asp 2) as β-secretase. Mol. Cell. Neurosci. 14, 419–427, 1999. 5. Lin, X. et al. Human aspartic protease memapsin 2 cleaves the β-secretase site of β-amyloid precursor protein. Proc. Natl. Acad. Sci. USA 97, 1456–1460, 2000. 6. Selkoe, D.J. Translating cell biology into therapeutic advances in Alzheimer’s disease. Nature 399A, 23–31, 1999. 7. Mullan, M. et al. A locus for familial early-onset Alzheimer’s disease on the long arm of chromosome 14, proximal to the alpha 1-antichymotrypsin gene. Nature Genet. 2, 340–342, 1992. 8. Ghosh, A.K. et al. Design of potent inhibitors for human brain memapsin 2 (β-secretase). J. Am. Chem. Soc. 122, 3522–3523, 2000. 9. Turner, R.T., III et al. Subsite specificity of memapsin 2 (β-secretase): implications for inhibitor design. Biochemistry 40, 10001–10006, 2001. 10. Hong, L. et al. Structure of the protease domain of memapsin 2 (β-secretase) complexed with inhibitor. Science 290, 150–153., 2000. 11. Hong, L. et al. Crystal structure of memapsin 2 (β-secretase) in complex with an inhibitor OM00-3. Biochemistry 41, 10963–10967, 2002. 12. Ermolieff, J. et al. Proteolytic activation of recombinant pro-memapsin 2 (pro-βsecretase) studied with new fluorogenic substrates. Biochemistry 39, 12450–12456, 2000. 13. Ermolieff, J., Lin, X., and Tang, J. Kinetic properties of saquinavir-resistant mutants of human immunodeficiency virus type 1 protease and their implications in drug resistance in vivo. Biochemistry 36, 12364–12370, 1997. 14. Co, E. et al. Proteolytic processing mechanisms of a miniprecursor of the aspartic protease of human immunodeficiency virus type I. Biochemistry 33, 1248–1254, 1994. 15. Liu, Y., Kati, W., Chen, C.-M., Tripathi, R., Molla, A., and Kohlbrenner, W. Use of a fluorescence plate reader for measuring kinetic parameters with inner filter effect correction. Anal. Biochem. 267, 331–335, 1999. 16. Ghosh, A.K., Hong, L., and Tang, J. β-Secretase as a therapeutic target for inhibitor drugs. Curr. Med. Chem. 9, 1135–1144, 2002. 17. Ghosh, A.K. et al. Structure-based design: potent inhibitors of human brain memapsin 2 (β-secretase). J. Med. Chem. 44, 2865–2868, 2001. 18. Bieth, J. Some kinetic consequences of the tight binding of protein-proteinase inhibitors to proteolytic enzymes and their application to the determination of dissociation constants, in Bayer Symposium V, Proteinase Inhibitors: Proceedings of the 2nd International Research Conference, Fritsch, H., Tschesche, H., and Greene, L.J., Eds., Springer-Verlag, Berlin, 1974, p. 463. 19. Leatherbarrow, R.J. GraFit Version 3.0, Erithacus Software Ltd., Staines, U.K., 1990. 20. Tomasselli, A.G. et al. Substrate analogue inhibition and active site titration of purified recombinant HIV-1 protease. Biochemistry 29, 264–269, 1990. 21. Marciniszyn, J., Jr., Hartsuck, J.A., and Tang, J. Mode of inhibition of acid proteases by pepstatin. J. Biol. Chem. 251, 7088–7093, 1976.

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5

Assays for Amyloid Precursor Protein γ-Secretase Activity William A. Campbell, Michael S. Wolfe, and Weiming Xia

CONTENTS Abstract 5.1 Introduction 5.2 Main Scheme of Approaches 5.3 Method 5.3.1 Assaying γ-Secretase Activity in Living Cells 5.3.2 Subcellular Fractionation of Membrane Vesicles 5.3.3 In Vitro γ-Secretase Activity Assay Using Endogenous Substrate 5.3.4 Determining pH Dependence of γ-Secretase Activity Using Vesicles 5.3.5 Protease Inhibitor Profiling of γ-Secretase Activity Using Fractions 5.3.6 Cell Membrane Preparation for Exogenous Substrate Assay 5.3.6.1 Buffers 5.3.7 M2 Flag Purification of E. coli-Generated γ-Secretase Substrates 5.3.7.1 Buffers 5.3.8 In Vitro γ-Secretase Activity Assay Using Exogenous Substrate 5.4 Discussion References

ABSTRACT γ-Secretase cleavage, mediated by a complex of presenilin (PS), nicastrin, PEN-2, and APH-1, is the final proteolytic step in generating amyloid beta (Aβ) protein and the Notch intracellular domain. Aβ and Notch are critical in the pathogenesis of Alzheimer’s disease (AD) and in development, respectively. In addition to cleaving amyloid precursor protein (APP) and Notch, γ-secretase also cleaves over a dozen additional type I transmembrane domain proteins. γ-Secretase activity can be measured in vivo by collecting conditioned media from tissue cultured cells and in vitro 0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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by incubating endogenous substrate with total microsomes or with Golgi/trans-Golgi network (TGN)-enriched microsomal vesicles or incubating recombinant substrate with solubilized membranes. For APP, Western blotting or enzyme-linked immunosorbent assay (ELISA) has been used to quantify the generation of Aβ40, Aβ42, and amyloid intracellular domain (AICD). These methods can be applied to study the γ-secretase cleavage of any γ-secretase substrate in a variety of experimental conditions.

5.1 INTRODUCTION Genetic and neuropathological studies suggest that processing of amyloid precursor protein (APP) to C99 and then to amyloid β protein (Aβ), the major component of destructive neuritic plaques found in brains of AD patients, plays an important role in the neuronal loss that leads to AD.1 The final proteolytic event in generating Aβ, which is mainly a 40- or 42-residue peptide, is accomplished through presenilin (PS)-dependent γ-secretase cleavage of the C99 peptide.1 In addition to APP, γ-secretase also cleaves over a dozen type I transmembrane proteins such as the Notch proteins involved in cell fate determination, ErbB4 tyrosine receptor kinase and cadherins.2–5 The growing list of γ-secretase substrates includes APP, Notch,2 E-cadherin and N-cadherin,5,6 ErbB4 tyrosine receptor kinase,4 CD44,7,8 Nectin1α,9 Delta and Jagged,10,11 LRP,12 DCC,13 APLP1 and APLP2,14–16 p75 neurotrophin receptor,17 Syndecan 3,18 glutamate receptor subunit 3,19 and colony stimulating factor 1.20 The biological significance of this cleavage is not clear in most cases, although one normal function may be to release intracellular domains that regulate gene transcription in the nucleus, in at least some cases. While absolute identification of the catalytic component of γ-secretase activity has been elusive, mounting evidence points to PS1 and PS2. Numerous studies have demonstrated that PS is necessary for γ-secretase cleavage and Aβ generation. For example, mutation of two critical aspartate residues in transmembrane (TM) domains 6 and 7 of PS1 or PS2 abolishes Aβ generation in cultured cells21–23 and in transgenic mice.24 PS knockout neurons do not produce any Aβ.25,26 PS1 and PS2 bind to the immediate substrates of γ-secretase, C99/C83, in the major sites of Aβ generation, i.e., Golgi/trans-Golgi network (TGN)-type vesicles.27 Finally, using aspartyl protease transition-state analogue γ-secretase inhibitors to probe the active site of the enzyme revealed that these inhibitors bind directly to PS N- and C-terminal fragments (NTF and CTF).28,29 PS and PS homologues also have nonclassic protease motifs conserved from bacteria to humans,30,31 and the sequence motifs of PS are similar to a signal peptide peptidase.32 Signal peptide peptidase forms a homodimer that is labeled by an active site-directed γ-secretase inhibitor, indicating that the active sites of signal peptide peptidase and PS/γ-secretase are similar.33 PS1 and PS2 are homologous eight transmembrane domain-spanning proteins that undergo constitutive endoproteolysis by an unknown enzyme termed presenilinase to generate functional stable heterodimers of NTF and CTF.34,35 Familial AD (FAD) mutations in PS1 or PS2 lead to increased production of the longer, more

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amyloidogenic version of Aβ, the 42-residue version, Aβ42.34,36–41 Knockout of PS1 in mice is embryonic lethal.42 Conditional knockout of PS1 in the brain results in deficits of long-term potentiation and cognition43 while restricted expression of PS1 in the brain in the background of PS1 knockout mice leads to skin tumorigenesis.44 Many studies have implicated the PS fragments, along with mature Nicastrin, APH-1 (anterior pharynx defective), and PEN-2 (presenilin enhancer), as the functional components of the γ-secretase complex. N-linked glycosylation of nicastrin in the Golgi apparatus is associated with its entry into the active γ-secretase complex, and this mature form interacts preferentially with the functional PS1 heterodimers.45–50 Down-regulation of APH-151,52 or PEN-251,53 by RNAi in cells is associated with reduced levels of PS1 NTF and CTF heterodimers and deficient γ-secretase function. Overexpressing APH-1 stabilizes the full-length (FL) PS1, whereas reducing PEN-2 decreases endoproteolytic processing of PS1.54–56 Many reports have shown that co-expression of PS1, Nicastrin, APH-1, and PEN-2 results in increased PS1 endoproteolysis and γ-secretase activity, both in mammalian cells54–59 and in yeast.60 To measure γ-secretase activity, cell-based assays using the endogenous C99 substrate were used initially. Living cells can be treated with γ-secretase inhibitors and γ-secretase activity can be measured by release of soluble Aβ into the tissue culture medium. Using total membrane vesicles isolated from tissue culture cells, γ-secretase was found to be pH-dependent and showed maximal activity at pH between 6.3 and 6.4.61 After separation of intact, fully functional membrane vesicles from cultured cells on discontinuous Iodixanol gradients, γ-secretase activity was predominantly localized to Golgi/TGN-rich vesicles.62 PS bound to the immediate substrates of γ-secretase, the C-terminal fragments of APP, in these Golgi/TGN-rich vesicles.27 Subsequently, γ-secretase activity was solubilized to partially characterize its activity using a recombinant substrate containing an initiating methionine, C99, and a Flag epitope (C100Flag). Anti-PS1 antibodies were found to immunoprecipitate γ-secretase activity from these solubilized membranes.63 Next, a Notch-based substrate, N100Flag, was created using an N-terminal methionine, 99 residues of the Notch1 sequence beginning from the ligand-dependent S2 cleavage site, and a C-terminal Flag sequence.64 The C100Flag and N100Flag substrates were then used to further characterize the γ-secretase activity that cleaves APP and Notch.64,65 With the help of these substrates and the detergent-dependent assay, the presenilin–γ-secretase complex was isolated from solubilized membrane preparations in an activitydependent manner using an immobilized active site-directed inhibitor, and the complex was found to contain Nicastrin and C83.64 A comparison of C100Flag and N100Flag proteolysis suggested that the responsible proteases are identical, with each substrate preventing cleavage of the other, and both substrates being cleaved at two distinct regions in the transmembrane domain.65

5.2 MAIN SCHEME OF APPROACHES Characterization of γ-secretase activity can be obtained by cellular assays of γ-secretase activity and by detergent solubilization of γ-secretase. The assays described present reliable and reproducible methods to measure the cleavage of the C99

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fragment of APP to generate Aβ40, Aβ42, and AICD, and should be applicable to measure the γ-secretase cleavage of any γ-secretase substrate. This protocol contains the procedures for analyzing γ-secretase activity in living cells in vitro using endogenous substrate in total membrane microsomes and in Golgi/TGN-rich fractions and in vitro using solubilized membranes and E. coligenerated recombinant γ-secretase substrates. The first method describes measuring Aβ secreted from living tissue culture cells. The second method describes the subcellular fractionation of membrane vesicles on discontinuous Iodixanol gradients to separate intact, functional ER-rich vesicles from Golgi/TGN-rich vesicles. In the third method, γ-secretase activity is measured in the Golgi/TGN-rich fractions. In the fourth method, the optimal pH for γ-secretase activity is determined using total membrane vesicles. The next method describes protease inhibitor profiling to discover the protease class of γ-secretase. The sixth method describes the preparation of solubilized cell membranes that retain functional γ-secretase enzymes. The seventh method describes the use of an M2 anti-Flag immunoaffinity isolation procedure to purify the γ-secretase substrates from E. coli. The last method details the γ-secretase activity assay using recombinant substrates. An outline of this scheme is presented in Figure 5.1. Subcellular fractionation of membrane organelles has been successfully used to measure γ-secretase activity in Golgi/TGN-rich vesicles.27,57,62,66,67 Solubilized γ-secretase has been used successfully to measure the cleavage of recombinant substrates C100Flag and N100Flag.63–65,68 Measuring secreted A β in living cells (Method 1)

Isolation of total microsomal membranes (Method 2) Endogenous substrate

Exogenous substrate

Subcellular fractionation (Method 2)

Cell membrane preparation (Method 6)

γ−secretase activity in Golgi/TGN-rich fractions Purification of substrates

(Method 3)

(Method 7)

pH dependence (Method 4)

Protease inhibitor profiling

γ−secretase activity in solubilized membranes

(Method 5)

(Method 8)

Western blot and/or ELISA

FIGURE 5.1 Methods discussed in this chapter.

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5.3 METHOD 5.3.1 ASSAYING γ-SECRETASE ACTIVITY

IN

LIVING CELLS

The first method to measure γ-secretase activity presented here is accomplished by collecting conditioned media from cultured cells treated either with a vehicle control or a γ-secretase inhibitor. Treating living cells with a γ-secretase inhibitor causes an increase in the immediate substrates of γ-secretase, the APP CTF’s C99 and C83, and decreases the total amount of Aβ secreted into the tissue culture medium (Figure 5.2). 1. Culture cells to confluence. 2. Make stock concentrations of a γ-secretase inhibitor in 100% dimethyl sulfoxide (DMSO). 3. Dilute the γ-secretase inhibitor or vehicle (DMSO) into the tissue culture media to achieve the desired final concentration in 1% DMSO. 4. After a 4-hr incubation, collect the conditioned medium and centrifuge it at 10,000 × g for 5 min to pellet cells or cell debris. 5. Remove the supernatant and store it at –80˚C until analysis by ELISA. 6. The cells can be collected, lysed and subjected to immunoprecipitation with APP polyclonal antibody C7 followed by Western blot analysis with a. 14 0

10

20

30

40

50

Dose (uM)

Relative Aβ Total

b.

1.0 0.9 0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0

n=8

0

7.5 10 12.5 15 17.5 20 22.5 25

30 40

50

Dose (uM)

FIGURE 5.2 Treatment of living cells with a γ-secretase inhibitor increases APP CTFs (C-terminal fragments) and decreases the amount of Aβ secreted into the media. (a) Chinese hamster ovary cells overexpressing wild-type (wt) human APP were treated with increasing concentrations (µM) of γ-secretase inhibitor compound 1,73,74 and cell lysates were immunoprecipitated with APP polyclonal antibody C7 followed by Western blotting with APP monoclonal antibody 13G8 to visualize the APP C terminal fragments (i.e., γ-secretase substrates). (b) Aβ levels in the conditioned media of cells treated with increasing doses of compound 1 were determined by ELISA. Aβ levels in each experiment were normalized to mock-treated (1% DMSO) samples and averaged (n = 8). (Reprinted from Xia, W. et al. Neurobiol. Dis. 2000, 7, 673–681. With permission.)

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APP monoclonal antibody 13G8 to visualize the APP C-terminal fragments (i.e., γ-secretase substrates). Polyclonal antibody C769 and monoclonal antibody 13G870 (gift of P. Seubert and D. Schenk) are directed against APP732–751 (APP751 numbering).

5.3.2 SUBCELLULAR FRACTIONATION

OF

MEMBRANE VESICLES

The fractionation employs discontinuous Iodixanol gradients that are used because they effectively separate ER- from Golgi/TGN-rich vesicles in a way that preserves vesicle structure and function, allowing detection of γ-secretase activity upon incubation of Golgi/TGN-rich vesicles at 37˚C.62,71 The viscosity of the gradient medium is a major determinant of the sedimentation rate. The osmotic activity of the medium is also important because subcellular organelles are osmotically sensitive. Thus, although sucrose, glycerol, and Ficoll are widely used for gradient fractionation of cellular membranes, they are not ideal in osmolality and viscosity. The main advantage of using an Iodixanol gradient is that osmolality and viscosity remain relatively constant with changes in the density of the gradient. Under this mild iso-osmotic condition, each organelle can be isolated functionally intact, without loss of water, as the density of the gradient increases. 1. Culture five 15-cm dishes until confluent (~1 × 108 cells). 2. Detach the cells with 20 mM EDTA in PBS (8 mL/15-cm plate). 3. Pellet the cells by spinning for 5 min at 4˚C, 1000 rpm. The cell pellet can be frozen at –80˚C indefinitely. 4. Resuspend the cell pellet in 3 mL of cold homogenization buffer (0.25 M sucrose, 10 mM HEPES, 1 mM EDTA) with freshly added protease inhibitors. 5. Break open the cells with ten strokes of a Dounce homogenizer and pass the cells through a 27-gauge needle five times. 6. Pellet the nuclei and unbroken cells by centrifugation at 1500 × g for 10 min at 4˚C. Save the postnuclear supernatant. 7. Extract the pellet again by resuspending in 4 mL of homogenization buffer and centrifuge at 1500 × g for 10 min at 4˚C. Save the postnuclear supernatant. 8. Combine the two supernatants and centrifuge for 1 hr at 65,000 × g, 4˚C, to pellet total membrane vesicles. 9. For measuring γ-secretase activity in total vesicles, the vesicles are washed in 0.1 M sodium carbonate (pH 11.3) on ice to remove peripherally associated membrane proteins and centrifuged at 100,000 × g for 1 hr. The vesicle precipitate is then resuspended in incubation buffer (10 mM KOAc, 1.5 mM MgCl2). One portion is lysed with Laemmli sample buffer (10% SDS, 0.3 M Tris, 50% glycerol, 0.1% bromophenol blue, 10% β-mercaptoethanol) for Western blot or 2X guanidium HCl (1 M guanidium HCl, 2% NP-40, 2 mM EDTA) for ELISA for basal γ-secretase activity. The other portion is incubated at 37˚C for de novo γ-secretase activity (see below).

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10. For separation of ER- and Golgi/TGN-rich vesicles, the membrane vesicles are used to prepare subcellular fractions by resuspending the microsomal pellet in 800 µL of homogenization buffer on ice. Different percentages of Iodixanol, 5,59-[(2-hydroxy-1-3- propanediyl)-bis(acetylamino)]bis[N,N-9-bis(2,3-dihydroxypropyl-2,4,6-triiodo-1,3-benzenecarboxamide] are made by diluting with OptiPrep (Accurate Chemical, Westbury, NY, 60% Iodixanol). 11. A gradient stock solution of 50% Iodixanol is prepared by diluting in 0.25 M sucrose, 6 mM EDTA, 60 mM HEPES, pH 7.4, at a 5:1 ratio. Different densities of Iodixanol are established by diluting this stock with 0.25 M sucrose homogenization buffer. 12. The Iodixanol gradient is poured in 13 mL Beckman SW41 centrifuge tubes by careful underlayering, adding each layer under the first layer with a long needle, as follows: 1 mL 2.5% Iodixanol 2 mL 5% Iodixanol 2 mL 7.5% Iodixanol 2 mL 10% Iodixanol 0.5 mL 12.5% Iodixanol 2 mL 15% Iodixanol 0.5 mL 17.5% Iodixanol 0.5 mL 20% Iodixanol 0.3 mL 30% Iodixanol 13. Load the resuspended vesicles on top of the gradient and centrifuge in an SW41 rotor at 200,000 × g for 2.5 hr at 4˚C. Decrease the acceleration and deceleration rates of the centrifuge so the gradient is not disturbed. 14. Collect 12 fractions ~1 mL at a time by puncturing the bottom of the tube with a 22-gauge needle. Seal the top of the tube with Parafilm prior to puncturing to control the flow of the gradient from the tube. Store the fractions indefinitely at –80˚C.

5.3.3 IN VITRO γ-SECRETASE ACTIVITY ASSAY USING ENDOGENOUS SUBSTRATE The 12 fractions must be characterized by Western blotting with the ER-specific marker calnexin and the Golgi/TGN-specific marker syntaxin 6 or by measuring the activity of the Golgi-specific enzyme galactosyltransferase to determine which fractions are Golgi/TGN-enriched.62,66 Typically, the first three fractions contain calnexinpositive ER-rich vesicles, the fourth fraction contains a mixture of ER and Golgi/TGN vesicles, and fractions five through eight contain Golgi/TGN-rich vesicles that can be used for measuring γ-secretase activity.62 Instead of using each individual fraction, the first four fractions can be combined and considered ER-rich vesicles; fractions five through eight can be combined and considered Golgi/TGNrich vesicles (Figure 5.3a). These vesicles can now be incubated at 37˚C for measurement of de novo γ-secretase activity, for example, by Aβ-specific ELISA or Western blotting for AICD after incubation.

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3.0

30

60

90

30

60

90

0.0

15 30

2

1

0

Golgi/TGN

16

120

1.0

d. min

120

15 15

90 120 120 PS1 cell lysate

5

10

5

10

3 3

60

2 2

1

Golgi/TGN

2.0

10

c.

0.0

1

< Syntaxin6

3 5

36

1.0 0

] Calnexin

Golgi/TGN 2.0

0.5

98

3.0

0

/TGN

0.5

b.

Golgi ER

Relative A β40Level

a.

Relative A β42Level

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6

FIGURE 5.3 Generation of Aβx-40, Aβx-42, and AICD in Golgi/TGN-rich microsome fractions. (a) Microsomes isolated from Chinese hamster ovary cells stably expressing wt APP and wt PS1 (PS1wt-1) were fractionated on discontinuous Iodixanol gradients. The first four 1-mL fractions were combined, and an aliquot was lysed and run in lanes 1 and 2. The next four 1-mL fractions were combined, and an aliquot was lysed and run in lanes 3 and 4. ER-enriched microsomes and Golgi/TGN-rich microsomes were lysed and probed by Western blot for the ER marker calnexin and the Golgi/TGN marker syntaxin 6. The first four fractions were enriched in the ER marker calnexin, while the next four fractions were enriched in the Golgi/TGN marker syntaxin 6. (b) and (c) The Golgi/TGN-enriched vesicles were lysed at time zero or incubated at 37˚C for the indicated times and then lysed. The lysed vesicles were then subjected to ELISA for measurement of de novo Aβx-40 and Aβx-42 generation. Aβ levels in each experiment were normalized to basal levels (denoted by dotted line) obtained at time 0. Error bars represent the standard error of the mean in all figures. (d) Golgi/TGN-rich microsomes were incubated at 37˚C up to 2 hr, lysed and probed by Western blot with polyclonal antibody R57 that readily detects AICD. Duplicate samples incubated at 37˚C for 120 min were presented. (Reprinted from Campbell, W.A. et al. J. Neurochem. 2003, 85, 1563–1574. With permission.)

1. On ice, thaw the fractions found to be Golgi/TGN-rich. If adding γ-secretase inhibitors or a vehicle control, add them to each well or tube at room temperature before adding the fractions on ice. Typically these reactions are run in a 96-well polymerase chain reaction (PCR) plate.

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2. Gently invert the Golgi/TGN-rich fractions and aliquot 50 µL into each cold tube or well for 4˚C and 37˚C samples. To the control 4˚C aliquot, immediately add 25 µL of 3X Laemmli sample buffer for determination of basal AICD levels, for example. If the goal is to measure Aβ by ELISA, then add 50 µL of 2X guanidinium HCl. 3. If adding γ-secretase inhibitors or DMSO, make sure that they are in solution and/or mixed well by heating the samples at 37˚C for 2 min and vortexing. This aliquot is incubated at 37˚C for 2 hr for determination of de novo Aβ or AICD generation. The reaction is stopped by adding 25 µL of 3X Laemmli sample buffer or 50 µL of 2X guanidinium HCl followed by vortexing. 4. AICD levels in both aliquots can be probed by Western blot. Densitometry using the AlphaEase™ software package (Alpha Innotech, San Leandro, CA) is used to quantify protein levels from at least three independent blots, and the level of newly generated AICD is calculated by subtracting the amount of AICD observed at 4˚C from that obtained at 37˚C. 5. For determination of de novo Aβ generation in vitro by ELISA, the newly generated Aβ levels are calculated by subtracting the Aβ level observed at 4˚C from that obtained at 37˚C. 6. For inhibition experiments with γ-secretase inhibitors, the mean levels of newly generated AICD or Aβ in the absence of inhibitor (H2O, DMSO or methanol) is used as the denominator (100%), and the relative percentage is obtained by comparing AICD or Aβ levels in the presence of inhibitor against this denominator in each experiment. Negative values reflect reduced levels after incubation. Results using this method in a time course experiment are presented in Figure 5.3b through Figure 5.3d. Generation of endogenous Aβx-40 and Aβx-42 as measured by ELISA and generation of endogenous AICD as measured by Western blot all begin approximately 10 min after incubation of Golgi/TGN-enriched vesicles at 37˚C.

5.3.4 DETERMINING PH DEPENDENCE USING VESICLES

OF

γ-SECRETASE ACTIVITY

This method has been used to show that the optimal pH for γ-secretase activity is 6.3 to 6.4 (Figure 5.4).61 Sodium citrate is added to total membrane vesicles to achieve a final pH gradient of 5.3 to 7.6. After removal of all cytosol and extensive washing of these microsomal vesicles, not much specific ionic exchange should occur among these membranes. The pH in the suspension should closely reflect the actual pH inside the vesicles. 1. Isolate total microsomal vesicles as described in method 5.3.2. 2. Resuspend microsomal vesicles in 5 mL incubation buffer (10 mM KOAc, 1.5 mM MgCl2) plus 50 mM sodium citrate, pH 5.6. 3. Titrate the pH by adding a battery of stock solutions of 1M sodium citrate with increasing pH (from 4.5 to 7.4) to isolated microsomes in incubation

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1.2 Aβ

a.

CHO

1.0

Relative

0.8 0.6 0.4 0.2 5.3 5.4 5.5 5.6 5.7 5.8 5.9 6.0 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9 7.0 7.1 7.2 7.3 7.4 7.5 7.6

0

1.2

b. Aβ

HEK293 1.0

Relative

0.8 0.6 0.4 0.2 7.4 7.5 7.6

5.3 5.4 5.5 5.6 5.7 5.8 5.9 6.0 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9 7.0 7.1 7.2 7.3

0

pH

FIGURE 5.4 pH dependence of Aβ generation in isolated microsomal vesicles. Stock solutions of sodium citrate with different pH levels were added to isolated microsomal vesicles from PS1WT-1 (a) or 293695SW (b) cells. Aliquots were incubated at various pH levels, and de novo Aβ generation was measured by ELISA. Relative Aβ levels generated at each pH were normalized with respect to the highest value (averages shown as x, with standard errors indicated by bars; PS1WT-1, n = 5; 293695SW, n = 6). A third-order polynomial curve fitting revealed a bell curve with the peak at pH 6.3 to 6.4. (Reprinted from Xia, W. et al. Neurobiol. Dis. 2000, 7, 673–681. With permission.)

buffers. This results in a concentration of 50 mM sodium citrate with the final pH ranging from 5.3 to 7.6, as measured by a microprobe. 4. For in vitro Aβ generation, total isolated microsomes in incubation buffer are divided into two aliquots. One aliquot is used for determination of basal Aβ levels by adding an equal volume of stop solution (2X guanidinium HCl) and freezing at –80˚C. 5. The other aliquot is incubated at 37˚C for 4 hr followed by addition of an equal volume of stop solution. 6. Aβ levels are determined in both aliquots by ELISA, and the newly generated Aβ levels are calculated by subtracting the Aβ level generated at –80˚C from that at 37˚C.

5.3.5 PROTEASE INHIBITOR PROFILING USING FRACTIONS

OF

γ-SECRETASE ACTIVITY

This method can be used to determine whether any drug of choice or which class of protease inhibitor can inhibit γ-secretase activity. For pharmacological inhibition

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of γ-secretase, gradient fractions are incubated with each protease inhibitor or γ-secretase inhibitor at the desired concentration or with DMSO, methanol, or H2O vehicle alone for 2 hr at 37˚C. 1. On ice, thaw the fractions found to be Golgi/TGN-rich. These reactions can easily be run in a 96-well PCR plate testing three concentrations of a particular inhibitor over three orders of magnitude. Add the protease inhibitor to a final concentration within the effective range of its activity to each well or tube at room temperature before adding the fractions on ice. 2. The four classes of protease inhibitors with the names of the inhibitors and their reported effective concentrations in parentheses are as follows: aspartyl (pepstatin A, 1 µM), metallo (1,10-phenathroline, 1 to 10 mM; EDTA, 1 to 10 mM; phosphoramidon, 1 to 10 µM), serine (Pefabloc, 0.4 to 4.0 mM; PMSF, 0.1 to 1.0 mM), and serine/cysteine (leupeptin, 10 to 100 µM).72 3. Gently invert the Golgi/TGN-rich fractions and aliquot 50 µL into each well on ice for 4 and 37˚C samples. To the control 4˚C aliquot, immediately add 25 µL of 3X Laemmli sample buffer for determination of basal AICD levels, for example. If the goal is to measure Aβ by ELISA then add 50 µL of 2X guanidinium HCl. 4. Ensure that the protease inhibitor is in solution by vortexing well, followed by heating the samples at 37˚C for 2 min, vortexing again, and incubating at 37˚C for 2 hr. 5. Fractions are then lysed and subjected to Western analysis or ELISA.

5.3.6 CELL MEMBRANE PREPARATION SUBSTRATE ASSAY

FOR

EXOGENOUS

Total membranes are prepared from a large batch of cells (e.g., Jurkat or HeLa) for measuring endogenous γ-secretase activity. It is important to remember that in order to maintain functional γ-secretase activity the membranes must be solubilized in a correct detergent that will not disrupt the γ-secretase complex, e.g., CHAPSO (Soltec Ventures, Beverly, MA). 1. Collect and pellet cells. For cells grown in suspension (such as Jurkat or HeLa cells), collect 6.25 × 107 cells per microsome preparation. A volume of 2.5 L of HeLa cells at 1 × 106 cells/mL will yield “40x” microsomes. For adherent cells, collect cells from five 15-cm dishes per microsome preparation. Centrifuge at ~3000 × g for 10 min. Cell pellets can be stored at –80˚C until needed. This is defined as a 1x microsome preparation. 2. Add 30 mL of MES buffer (with 100x protease inhibitors added freshly) to 120x of microsomes and fully resuspend the cells. 3. Pass the cells once through a French pressure cell at greater than 1000 psi. 4. Pellet the nuclei and unbroken cells by centrifuging the samples at 3000 × g for 10 min (4550 rpm in a Sorvall SA600 rotor). Save the postnuclear supernatant.

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5. Pellet total membranes by centrifuging the supernatant at 100,000 × g for 1 hr (30,000 rpm in SW55 rotor; 25,000 rpm in SW41 rotor). Discard the supernatant. 6. Microsome pellets as 10x or 20x pellets can be stored at –80˚C until needed. 7. Resuspend each 10x microsome pellet in 0.5 mL of ice cold bicarbonate buffer. Pipet up and down at least 30 times. Incubate 20 min at 4˚C. 8. Pellet the washed membranes by centrifuging at 100,000 × g for 1 hr at 4˚C and discard the supernatant. 9. Resuspend the membranes by adding 0.5 mL of 1% CHAPSO lysis buffer to a bicarbonate-washed 10x microsome pellet. Pipet up and down at least 30 times. Vortex briefly and incubate for 1 hr at 4˚C. 10. Pellet the insoluble material by centrifugation at 100,000 × g for 1 hr at 4˚C. Discard the pellet. Dilute the supernatant 1:3 with HEPES buffer (final CHAPSO concentration is 0.25%). This diluted lysate is defined as solubilized γ-secretase. Measure the protein concentration with BCA reagent. The concentration should be ~0.10 to 0.25 mg/mL. Samples can be aliquoted and stored at –80˚C. 5.3.6.1 Buffers MES Buffer: 50 mM MES, pH 6.0, 150 mM NaCl, 5 mM MgCl2, 5 mM CaCl2, 100X complete protease inhibitor cocktail (Roche) added freshly Sodium Bicarbonate buffer: 0.1 M NaHCO3, pH 11.3 CHAPSO lysis buffer: 1% CHAPSO, HEPES buffer HEPES buffer: 50 mM HEPES, pH 7.0, 150 mM NaCl, 5 mM MgCl2, 5 mM CaCl2

5.3.7 M2 FLAG PURIFICATION γ-SECRETASE SUBSTRATES

OF

E.

COLI-GENERATED

To examine γ-secretase activity in vitro without the complication of cellular material, a recombinant substrate such as C100Flag can be generated to measure activity. The substrate is expressed in E. coli and purified using an M2 anti-Flag immunoaffinity resin. 1. Inoculate a 5-mL Luria Broth (LB) culture (with 100 µg/mL ampicillin) of BL21 (DE3)-transformed E. coli by scraping a small amount of glycerol culture, shaking at 37˚C throughout the day and storing at 4˚C overnight. 2. Inoculate the small culture into 250 mL LB medium and grow at 37˚C until OD600 = 1.0. 3. Induce protein expression by adding 1 mM IPTG (59.5 mg/250 mL) and 100 µg/mL ampicillin into the cultures and return to 37˚C for 2 hr. 4. Collect the cell pellet by centrifuging the cells at 3000 × g for 10 min. 5. Lyse 125 mL of cell pellet in 10 mL of 1% Triton X-100 lysis buffer with protease inhibitors and pass through the French press twice at pressure above 1000 psi.

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6. Spin down the insoluble pellet by centrifugation at 3000 × g for 10 min. 7. Bind to the M2 anti-Flag resin for 2 hr by adding 2 mL of M2 anti-Flag resin (4 mL of slurry) and rock for 2 hr at room temperature. 8. Pour into a 2-mL column and wash the column with lysis buffer three times with two column volumes (12 mL total). 9. Elute the substrate with acidic glycine, collecting five fractions of 2 mL each. Re-equilibrate the column immediately with 10 mL PBS, then exchange into storage buffer. Store the column at 4˚C until reuse. The column can be used up to three times. Analyze each fraction on a 4 to 30% Tris-glycine gel and stain with Coomassie blue (Gelcode Blue). 5.3.7.1 Buffers Lysis buffer: 10 mM Tris, pH 7.0, 150 mM NaCl, 1% Triton X-100, 1X bacterial protease inhibitor (Sigma, St. Louis, MO) Elution buffer: 1% NP-40, 100 mM glycine, pH 2.7 Storage buffer: 50% glycerol, phosphate buffered saline, 0.02% Na azide

5.3.8 IN VITRO γ-SECRETASE ACTIVITY ASSAY USING EXOGENOUS SUBSTRATE To measure γ-secretase activity in the solubilized membrane preparation, it is relatively straightforward to combine the substrate and the membranes. Since the C100Flag protein tends to form high molecular weight aggregates, perhaps due to the presence of the Aβ sequence, SDS must be added to the substrate to disaggregate it. The assay must be performed in a detergent such as CHAPSO that is compatible with γ-secretase activity. The addition of phosphatidylcholine (PC) and phosphatidylethanolamine (PE) at certain concentrations can augment γ-secretase activity.65 After incubation of the membranes at 37˚C, the three predominant cleavages made by γ-secretase can be detected by either Western blot or ELISA using the appropriate antibodies. 1. Thaw bicarbonate-washed, detergent-solubilized HeLa cell membranes (0.25% CHAPSO HEPES) and substrate (C100Flag) on ice. 2. Disaggregate the substrate by heating in SDS. Add SDS to 0.5% final concentration, then heat for 5 min at 65˚C. Centrifuge at 14,000 rpm for 2 min to pellet the insoluble C100Flag aggregates. 3. Aliquot into a 96-well plate or microfuge tubes. For each assay, aliquot 50 µL of solubilized HeLa cell membranes at 0.150 mg/mL and add 2.5 µL of PE (5 mg/mL in 1% CHAPSO–HEPES) and 5 µL of PC (10 mg/mL in 1% CHAPSO–HEPES) to a final concentration of 0.025% PE and 0.100% PC. Now add 1 µL of C100FLAG. For time 0, add 15 µL of 4X Laemmli sample buffer prior to adding the substrate. If the sample will be analyzed by Aβ ELISA, stop with 10 µL of 2.5% SDS. 4. Mix briefly by vortexing, cover with a 96-well plate sealer to prevent evaporation and incubate for 4 hr at 37˚C.

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5. Stop the reaction by adding 15 µL of 4X Laemmli sample buffer or 10 µL of 2.5% SDS solution and mix. 6. Analyze by M2 anti-Flag western blot or Aβ ELISA. Load 20 µL of sample onto an Invitrogen 4 to 20% Tris-glycine 10- or 12-well gel. Transfer and western blot with M2 anti-Flag antibody (Sigma) using standard protocols. Alternatively, analyze the C100Flag reaction for Aβx-40 and/or Aβx-42 proteolytic products by ELISA. 7. Important points to consider about the enzyme: It is stable at 4˚C for at least 1 week and survives three freeze–thaws, but detergents such as NP-40, Triton X-100 and SDS disrupt activity. Optimal pH is slightly acidic to neutral (6.5 to 7.0) and CHAPSO, CHAPS, and Big CHAP detergents are critical for retaining activity, preferably below the critical micelle concentration (0.25% for CHAPSO).

5.4 DISCUSSION This chapter has described assays for measuring γ-secretase activity in whole cells, total membranes, Golgi/TGN-enriched membrane vesicles from cultured cells, and in a solubilized membrane preparation using recombinant substrates. We also discussed methods for determining the optimal pH of γ-secretase activity and using protease inhibitor profiling for γ-secretase activity. While we have described these assays to examine the γ-secretase-mediated cleavage of APP, certainly these protocols can be adapted to examine the cleavage of any γ-secretase substrate. These methods have already been applied to Notch cleavage, for example.64,65 In any case, it is always necessary to include controls. For the Golgi/TGN-enriched vesicles, the collected fractions must be characterized to confirm that they are Golgi/TGNenriched and a γ-secretase inhibitor should be included during incubation as a control. For the substrate assays, it is also important to disaggregate the substrate and include a γ-secretase inhibitor as a control. The amounts of PE and PC added to the assay may need to be optimized for each substrate. In summary, the approaches described in this chapter provide methods to analyze γ-secretase activity in whole cells, in vesicles isolated from cells, or in solubilized membranes. In this way, the γ-secretase cleavage of each substrate as well as the inherent characteristics of the enzyme itself can be analyzed.

REFERENCES 1. Selkoe, D.J. and Podlisny, M.B. Deciphering the genetic basis of Alzheimer’s disease. Annu. Rev. Genomics Hum. Genet. 3, 67, 2002. 2. De Strooper, B. et al. A presenilin-1-dependent γ-secretase-like protease mediates release of Notch intracellular domain. Nature 398, 518, 1999. 3. Ni, C.Y. et al. γ-Secretase cleavage and nuclear localization of ErbB-4 receptor tyrosine kinase. Science 294, 2179, 2001. 4. Lee, H.J. et al. Presenilin-dependent γ-secretase-like intramembrane cleavage of ErbB4. J. Biol. Chem. 277, 6318, 2002.

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5. Marambaud, P. et al. A presenilin-1/γ-secretase cleavage releases the E-cadherin intracellular domain and regulates disassembly of adherens junctions. EMBO J. 21, 1948, 2002. 6. Marambaud, P. et al. A CBP binding transcriptional repressor produced by the PS1/εcleavage of N-cadherin is inhibited by PS1 FAD mutations. Cell 114, 635, 2003. 7. Lammich, S. et al. Presenilin-dependent intramembrane proteolysis of CD44 leads to the liberation of its intracellular domain and the secretion of an Aβ-like peptide. J. Biol. Chem. 277, 44754, 2002. 8. Murakami, D. et al. Presenilin-dependent γ-secretase activity mediates the intramembranous cleavage of CD44. Oncogene 22, 1511, 2003. 9. Kim, D.Y., Ingano, L.A., and Kovacs, D.M. Nectin-1α, an immunoglobulin-like receptor involved in the formation of synapses, is a substrate for presenilin/γ-secretase-like cleavage. J. Biol. Chem. 277, 49976, 2002. 10. Ikeuchi, T. and Sisodia, S.S. The Notch ligands, Delta1 and Jagged2, are substrates for presenilin-dependent “γ-secretase” cleavage. J. Biol. Chem. 278, 7751, 2003. 11. LaVoie, M.J. and Selkoe, D.J. The Notch ligands, Jagged and Delta, are sequentially processed by α-secretase and presenilin/γ-secretase and release signaling fragments. J. Biol. Chem. 278, 34427, 2003. 12. May, P., Reddy, Y.K., and Herz, J. Proteolytic processing of low density lipoprotein receptor-related protein mediates regulated release of its intracellular domain. J. Biol. Chem. 277, 18736, 2002. 13. Taniguchi, Y., Kim, S.H., and Sisodia, S.S. Presenilin-dependent “γ-secretase” processing of deleted in colorectal cancer (DCC). J. Biol. Chem. 278, 30425, 2003. 14. Walsh, D.M. et al. γ-Secretase cleavage and binding to FE65 regulate the nuclear translocation of the intracellular C-terminal domain (ICD) of the APP family of proteins. Biochemistry 42, 6664, 2003. 15. Scheinfeld, M.H. et al. Processing of beta-amyloid precursor-like protein-1 and -2 by γ-secretase regulates transcription. J. Biol. Chem. 277, 44195, 2002. 16. Eggert, S. et al. The proteolytic processing of the amyloid precursor protein gene family members APLP-1 and APLP-2 involves α-, β-, γ-, and ε-like cleavages. Modulation of APLP-1 processing by N-glycosylation. J. Biol. Chem. 279, 18146, 2004. 17. Kanning, K.C. et al. Proteolytic processing of the p75 neurotrophin receptor and two homologs generates C-terminal fragments with signaling capability. J. Neurosci. 23, 5425, 2003. 18. Schulz, J.G. et al. Syndecan 3 intramembrane proteolysis is presenilin/γ-secretasedependent and modulates cytosolic signaling. J. Biol. Chem. 278, 48651, 2003. 19. Meyer, E.L. et al. Glutamate receptor subunit 3 is modified by site-specific limited proteolysis including cleavage by γ-secretase. J. Biol. Chem. 278, 23786, 2003. 20. Wilhelmsen, K. and van der Geer, P. Phorbol 12-myristate 13-acetate-induced release of the colony-stimulating factor 1 receptor cytoplasmic domain into the cytosol involves two separate cleavage events. Mol. Cell. Biol. 24, 454, 2004. 21. Wolfe, M.S. et al. Two transmembrane aspartates in presenilin-1 required for presenilin endoproteolysis and γ-secretase activity. Nature 398, 513, 1999. 22. Steiner, H. et al. A loss of function mutation of presenilin-2 interferes with amyloid β-peptide production and Notch signaling. J. Biol. Chem. 274, 28669, 1999. 23. Kimberly, W.T. et al. The transmembrane aspartates in presenilin 1 and 2 are obligatory for γ-secretase activity and amyloid β-protein generation. J. Biol. Chem. 275, 3173, 2000.

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24. Xia, X. et al. The aspartate-257 of presenilin 1 is indispensable for mouse development and production of β-amyloid peptides through β-catenin-independent mechanisms. Proc. Natl. Acad. Sci. USA 99, 8760, 2002. 25. De Strooper, B. et al. Deficiency of presenilin-1 inhibits the normal cleavage of amyloid precursor protein. Nature 391, 387, 1998. 26. Herreman, A. et al. Total inactivation of γ-secretase activity in presenilin-deficient embryonic stem cells. Nat. Cell Biol. 2, 461, 2000. 27. Xia, W. et al. Presenilin complexes with the C-terminal fragments of amyloid precursor protein at the sites of amyloid β-protein generation. Proc. Natl. Acad. Sci. USA 97, 9299, 2000. 28. Esler, W.P. et al. Transition-state analogue inhibitors of γ-secretase bind directly to Presenilin-1. Nat. Cell Biol. 2, 428, 2000. 29. Li, Y.-M. et al. Photoactivated γ-secretase inhibitors directed to the active site covalently label presenilin 1. Nature 405, 689, 2000. 30. Steiner, H. et al. Glycine 384 is required for presenilin-1 function and is conserved in bacterial polytopic aspartyl proteases. Nat. Cell Biol. 2, 848, 2000. 31. Ponting, C. et al. Identification of a novel family of presenilin homologues. Hum. Mol. Genet. 11, 1037, 2002. 32. Weihofen, A. et al. Identification of signal peptide peptidase, a presenilin-type aspartic protease. Science 296, 2215, 2002. 33. Nyborg, A.C. et al. Signal peptide peptidase forms a homodimer that is labeled by an active site directed γ-secretase inhibitor. J. Biol. Chem. 5, 5, 2004. 34. Borchelt, D.R. et al. Familial Alzheimer’s disease-linked presenilin 1 variants elevate Aβ1-42/1-40 ratio in vitro and in vivo. Neuron 17, 1005, 1996. 35. Thinakaran, G. et al. Endoprotreolysis of presenilin 1 and accumulation of processed derivatives in vivo. Neuron 17, 181, 1996. 36. Lemere, C.A. et al. The E280A presenilin 1 Alzheimer mutation produces increased Aβ42 deposition and severe cerebellar pathology. Nat. Med. 2, 1146, 1996. 37. Borchelt, D.R. et al. Accelerated amyloid deposition in the brains of transgenic mice coexpressing mutant presenilin 1 and amyloid precursor proteins. Neuron 19, 939, 1997. 38. Citron, M. et al. Mutant presenilins of Alzheimer’s disease increase production of 42-residue amyloid β-protein in both transfected cells and transgenic mice. Nat. Med. 3, 67, 1997. 39. Duff, K. et al. Increased amyloid-β42(43) in brains of mice expressing mutant presenilin 1. Nature 383, 710, 1996. 40. Petanceska, S.S. et al. Mutant presenilin 1 increases the levels of Alzheimer amyloid beta-peptide Aβ42 in late compartments of the constitutive secretory pathway. J. Neurochem. 74, 1878, 2000. 41. Xia, W. et al. Enhanced production and oligomerization of the 42-residue amyloid β-protein by Chinese hamster ovary cells stably expressing mutant presenilins. J. Biol. Chem. 272, 7977, 1997. 42. Shen, J. et al. Skeletal and CNS defects in presenilin-1-deficient mice. Cell 89, 629, 1997. 43. Yu, H. et al. APP processing and synaptic plasticity in presenilin-1 conditional knockout mice. Neuron 31, 713, 2001. 44. Xia, X. et al. Loss of presenilin 1 is associated with enhanced beta-catenin signaling and skin tumorigenesis. Proc. Natl. Acad. Sci. USA 98, 10863, 2001. 45. Arawaka, S. et al. The levels of mature glycosylated nicastrin are regulated and correlate with γ-secretase processing of amyloid β-precursor protein. J. Neurochem. 83, 1065, 2002.

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46. Kimberly, W.T. et al. Complex N-linked glycosylated nicastrin associates with active γ-secretase and undergoes tight cellular regulation. J. Biol. Chem. 277, 35113, 2002. 47. Leem, J.Y. et al. Presenilin 1 is required for maturation and cell surface accumulation of nicastrin. J. Biol. Chem. 277, 19236, 2002. 48. Tomita, T. et al. Complex N-glycosylated form of nicastrin is stabilized and selectively bound to presenilin fragments. FEBS Lett. 520, 117, 2002. 49. Yang, D.S. et al. Mature glycosylation and trafficking of nicastrin modulate its binding to presenilins. J. Biol. Chem. 277, 28135, 2002. 50. Chen, F. et al. Presenilin 1 and presenilin 2 have differential effects on the stability and maturation of nicastrin in mammalian brain. J. Biol. Chem. 278, 19974, 2003. 51. Francis, R. et al. Aph-1 and pen-2 are required for Notch pathway signaling, γ-secretase cleavage of βAPP and presenilin protein accumulation. Dev. Cell 3, 85, 2002. 52. Lee, S. et al. Mammalian APH-1 interacts with presenilin and nicastrin, and is required for intramembrane proteolysis of APP and Notch. J. Biol. Chem. 277, 45013, 2002. 53. Steiner, H. et al. PEN-2 is an integral component of the γ-secretase complex required for coordinated expression of presenilin and nicastrin. J. Biol. Chem. 277, 39062, 2002. 54. Hu, Y. & Fortini, M. Different cofactor activities in γ-secretase assembly: evidence for a nicastrin-Aph-1 subcomplex. J. Cell Biol. 161, 685, 2003. 55. Luo, W.J. et al. PEN-2 and APH-1 coordinately regulate proteolytic processing of presenilin 1. J. Biol. Chem. 278, 7850, 2003. 56. Takasugi, N. et al. The role of presenilin cofactors in the γ-secretase complex. Nature 422, 438, 2003. 57. Baulac, S. et al. Functional γ-secretase complex assembly in Golgi/trans-Golgi network: interactions among presenilin, nicastrin, Aph1, Pen-2 and γ-secretase substrates. Neurobiol. Dis. 14, 194, 2003. 58. De Strooper, B. Aph-1, Pen-2, and nicastrin with presenilin generate an active γ-secretase complex. Neuron 38, 9, 2003. 59. Kimberly, W. et al. γ-Secretase is a membrane protein complex comprised of presenilin, nicastrin, Aph-1, and Pen-2. Proc. Natl. Acad. Sci. USA 100, 6382, 2003. 60. Edbauer, D. et al. Reconstitution of γ-secretase activity. Nat. Cell Biol. 7, 7, 2003. 61. Xia, W. et al. FAD mutations in presenilin-1 or amyloid precursor protein decrease the efficacy of a γ-secretase inhibitor: a direct involvement of PS1 in the γ-secretase cleavage complex. Neurobiol. Dis. 7, 673, 2000. 62. Xia, W. et al. Presenilin 1 regulates the processing APP C-terminal fragments and the generation of amyloid β-protein in ER and Golgi. Biochemistry 37, 16465, 1998. 63. Li, Y.M. et al. Presenilin 1 is linked with γ-secretase activity in the detergent solubilized state. Proc. Natl. Acad. Sci. USA 97, 6138, 2000. 64. Esler, W.P. et al. Activity dependent isolation of the presenilin-γ-secretase complex reveals nicastrin and a γ substrate. Proc. Natl. Acad. Sci. USA 99, 2720, 2002. 65. Kimberly, W.T. et al. Notch and the amyloid precursor protein are cleaved by similar γ-secretase(s). Biochemistry 42, 137, 2003. 66. Campbell, W. et al. Endoproteolysis of presenilin in vitro: inhibition by γ-secretase inhibitors. Biochemistry 41, 3372, 2002. 67. Campbell, W.A. et al. Presenilin endoproteolysis mediated by an aspartyl protease activity pharmacologically distinct from γ-secretase. J. Neurochem. 85, 1563, 2003. 68. Xu, M. et al. γ-Secretase: characterization and implication for Alzheimer disease therapy. Neurobiol. Aging 23, 1023, 2002. 69. Podlisny, M.B., Tolan, D., and Selkoe, D.J. Homology of the amyloid β-protein precursor in monkey and human supports a primate model for β-amyloidosis in Alzheimer’s disease. Am. J. Pathol. 138, 1423, 1991.

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70. Oltersdorf, T. et al. The Alzheimer amyloid precursor protein: identification of a stable intermediate in the biosynthetic/degradative pathway. J. Biol. Chem. 265, 4492, 1990. 71. Zhang, J. et al. Subcellular distribution and turnover of presenilins in transfected cells. J. Biol. Chem. 273, 12436, 1998. 72. Beynon, R. and Salvesen, G., Proteolytic Enzymes: A Practical Approach. Beynon, R. and Bond, J., Eds., Oxford University Press, New York, 2001, p. 317. 73. Wolfe, M.S. et al. A substrate-based difluoro ketone selectively inhibits Alzheimer’s γ-secretase activity. J. Med. Chem. 41, 6, 1998. 74. Wolfe, M.S. et al. Peptidomimetic probes and molecular modeling suggest Alzheimer’s γ-secretase is an intramembrane-cleaving aspartyl protease. Biochem. 38, 4720, 1999.

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6

Cell-Free Reconstitution of β-Amyloid Production and Trafficking Dongming Cai, William J. Netzer, Feng Li, and Huaxi Xu

CONTENTS Abstract 6.1 Introduction 6.2 Methods 6.2.1 Pulse-Labeling of N2a Cells and Temperature Blocking of Nascent Protein Transport 6.2.2 Preparation of Permeabilized N2a Cells through Osmotic Shock and Mechanical Shear 6.2.3 Preparation of Cytosol for Cell-Free Reconstitution System 6.2.4 Preparation of Energy-Regenerating System for Cell-Free Reconstitution System 6.2.5 Aβ Generation in Cell-Free System Utilizing ERS in the Absenc of Cytosol 6.2.6 Detection of Aβ Production from Cell-Free System 6.2.7 Formation of Nascent Secretory Vesicles in Cell-Free System Supplemented with ERS and Cytosol 6.2.8 Detection of βAPP in Different Fractions through Immunoprecipitation 6.3 Discussion 6.3.1 ATP-Dependent Aβ Generation in Cell-Free Reconstitution System 6.3.2 Cytosol-Dependent βAPP Trafficking from TGN/ER in the Cell-Free Reconstitution System 6.3.3 Utilizing the Cell-Free Reconstitution System to Demonstrate PS1-Regulated Intracellular Trafficking and Surface Delivery of βAPP 6.3.4 Characterization of Vesicle and Membrane Fractions by Sucrose Gradient and Electron Microscopy Acknowledgments References 0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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ABSTRACT The production of β-amyloid is one aspect of the metabolism of βAPP. To achieve an effective understanding of this process, its must be considered in terms of both the enzymatic activities required for the proteolytic processing of βAPP and the trafficking of βAPP and its metabolites within the secretory pathway. Whole cells are often insufficient to elucidate the details of this process. We have therefore developed a cell-free reconstitution system for β-amyloid and βAPP production and trafficking. This experimental system has allowed us to study the effects of adding or removing salient cellular factors and to focus on βAPP processing as it occurs in various subcellular compartments.

6.1 INTRODUCTION Alzheimer’s disease (AD) is characterized by the excessive generation and accumulation of β-amyloid (Aβ) peptides. The amyloidogenic Aβ peptide is proteolytically derived from the β-amyloid precursor protein (βAPP) within the secretory pathway by distinct enzymatic activities known as β- and γ-secretase.1,2 Full-length βAPP is synthesized in the endoplasmic reticulum (ER) and transported through the Golgi apparatus. The major population of secreted Aβ peptides is generated within the trans-Golgi-network (TGN),3–5 also the major site of βAPP residence in neurons at steady state. βAPP can be transported in TGN-derived secretory vesicles to the cell surface if not first proteolyzed to Aβ or an intermediate metabolite. At the plasma membrane βAPP is either cleaved by the α-secretase activity to produce a soluble molecule, sβAPP,6 or alternatively, reinternalized within clathrincoated vesicles to an endosomal/lysosomal degradation pathway.7,8 The endocytic compartments have also been shown to contribute to Aβ generation.9,10 Thus, the distribution of βAPP between the TGN and cell surface has a direct influence upon the relative generation of sβAPP versus Aβ. This phenomenon makes delineation of the mechanisms responsible for regulating βAPP trafficking from the TGN/ER relevant to understanding the pathogenesis of AD. It has been suggested that AD pathology rests on the cellular processes that regulate βAPP metabolism and its localization within the secretory compartment. Specifically, it was demonstrated that Aβ 1-40 and various N terminal truncated Aβ variants (x-40) are produced largely in the TGN and packaged into post-TGN secretory vesicles. It was also demonstrated that Aβ1-42 (one of the most amyloidogenic forms) and Aβx-42 are produced in the TGN and are also packaged into post-TGN secretory vesicles.5 In addition, a fraction of the N-terminally truncated Aβ42, Aβx-42, can be generated and remain insoluble in the ER.5,11 These studies suggest that the accumulation of intracellular Aβ may be important in AD pathology and that stable intracellular pools of insoluble Aβ may exist. In addition, studies by our group suggest that the α-secretase activity is up-regulated by protein kinase C (PKC).12 This phenomenon was investigated by utilizing a cellfree system consisting of isolated TGN from PC12 cells, which allowed us to control

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the exposure of viable secretory membranes to cytosolic factors that could affect βAPP metabolism and vesicle trafficking.28 It was also demonstrated, using a similar experimental approach, that activation of PKA increases trafficking of βAPP from the TGN, but through a mechanism that is distinct from PKC.13 Recently, we reported that 17β-estradiol (17β-E2), a gonadal steroid hormone known to reduce Aβ formation both in cells and in AD transgenic mice,14,15 stimulates formation of vesicles containing βAPP from the TGN. Accelerated βAPP trafficking precludes maximal Aβ generation within the TGN.16 The cell-free system described herein has been used successfully to investigate the formation of nascent secretory vesicles containing various proteins such as prohormones, VSV G protein and βAPP, as well as proteolytic cleavage of βAPP in a variety of cells.5,17,18 This cell-fee reconstitution system has been well characterized, e.g., the integrity of Golgi stacks and nascent vesicles has been demonstrated by electron microscopy, and the intactness of the nascent vesicles has been shown by the resistance of luminal cargo proteins to proteinase K digestion.19 It is well known that trafficking of proteins to the cell surface from TGN/ER depends on the recruitment of cytosolic trafficking factors and the precise regulation of phospholipid composition.20,21 In addition, vesicle biogenesis is also dependent on the correct conformation or folding of the cargo molecule or the proper interactions between each component of a complex cargo. One of the best examples is that the transport of VSV G protein from the ER requires oligomerization of its monomers, a process of protein–protein interaction strictly dependent upon adenosine triphosphate (ATP) and temperature. Failure of proper oligomerization, by ATP depletion or incubation at nonpermissive temperatures severely impairs the exit of VSV G from the ER.22,23 Interestingly, our results also demonstrated an ATP requirement for Aβ formation at the γ-secretase cleavage step.3,24 Therefore, supplementation of cytosol and/or energy in the cell-free reconstitution system has been crucial for analyzing the metabolism and trafficking of βAPP through the secretory pathway. In addition, it has been well established that incubation of cells at 15 or 20oC leads to an accumulation of membrane and secretory proteins in the ER and TGN, respectively.17,25 These phenomena were first taken advantage of in cell-free systems that reconstituted prohormone cleavage and maturation which allow accumulation of prohormones in the respective compartments without being proteolyzed at the low temperatures.17 In general, small peptide hormones are synthesized as longer polyproteins that must be cleaved endoproteolytically at specific sites to yield the mature peptide hormones. Using a cell-free system along with temperature blocks allowed both the cellular sites of prohormone maturation and necessary cellular factors to be elucidated. Similar methods have been co-opted successfully for the study of βAPP processing. The objective of this chapter is to provide a thorough outline of experimental procedures for preparation of the cell-free reconstitution system and the analysis of Aβ production as well as intracellular trafficking of βAPP from the TGN and the ER in this system.

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6.2 METHODS 6.2.1 PULSE-LABELING OF N2A CELLS AND TEMPERATURE BLOCKING OF NASCENT PROTEIN TRANSPORT 1. Mouse neuroblastoma N2a cells overexpressing human βAPP variants are grown in culture media up to 80% confluent (one 100-mm plate is sufficient to prepare two cell-free reactions). 2. Wash cells on plate in 2 ml Dulbecco’s Modified Eagle Medium (DMEM) media without methionine. Then starve cells in 2 ml DMEM without methionine for 30 min at 37oC. 3. Cells are then labeled with [35S]methionine (500 µCi/ml) for 10 to 15 min at 37oC. 4. Wash cells with PBS (prewarmed or equilibrated to 20oC) three times. 5. Add 2 or 3 ml complete media equilibrated to 20oC and incubate the cells at 20˚C for 2 hr by floating plates in an undisturbed water bath. At this temperature, the transport of proteins, including βAPP, from the ER to the TGN is unimpaired. However, under these conditions, the egress of secretory vesicles from the TGN is blocked, thus allowing labeled fulllength APP (and other newly synthesized, labeled membrane proteins) to accumulate in the TGN.3,5,16 This experimental design is used when either TGN-specific vesicle biogenesis or βAPP metabolism is assayed. 6. Alternatively, to assay βAPP trafficking from the ER or APP metabolism in the ER, cells are labeled with [35S]-methionine for 4 hr at 15˚C to accumulate βAPP within the ER and block its transport to the TGN.

6.2.2 PREPARATION OF PERMEABILIZED N2A CELLS OSMOTIC SHOCK AND MECHANICAL SHEAR

THROUGH

For both TGN and ER temperature block experiments, cells are permeabilized after the 2- or 4-hr chase as follows. 1. Transfer plates to 4oC immediately. 2. Cells are incubated at 4oC in 3 ml “swelling buffer” (10 mM KCl, 10 mM HEPES, pH 7.2) for 10min. 3. The buffer is discarded and replaced with 1 ml of “breaking buffer” (90 mM KCl, 10 mM HEPES, pH7.2), after which the cells are broken by scraping vigorously with a rubber policeman. 4. The permeabilized cells are centrifuged at 800 × g for 5 min to remove cytosol, and washed in 5 ml of breaking buffer. The permeabilized cells must be washed and resuspended at least three times in order to remove cytosol and resuspended in five pellet volumes of breaking buffer. This should result in >95% cell breakage as evaluated by trypan blue staining. 5. Broken cells (cell-free system) are resuspended and incubated in a final volume of 300 µl containing 2.5 mM MgCl2, 0.5 µM CaCl2, and 110 mM KCl, an energy-regenerating system (ERS), with or without cytosol (15 µg proteins) prepared from N2a or other cells as needed.16,19 A protease

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inhibitor cocktail is added for vesicle budding experiments but not when Aβ production is assayed.

6.2.3 PREPARATION OF CYTOSOL FOR CELL-FREE RECONSTITUTION SYSTEM 1. Ten plates of N2a (or other desired) cells are cultured and collected in their media when grown up to 90% confluent. 2. Pellet cells down by centrifugation at 800 × g for 5 min. 3. Wash cell pellet with TEA buffer (10 mM triethanolamine, 140 mM KOAc, pH 7.2) followed by a wash in homogenization buffer (25 mM HEPES–KOH, pH 7.2, 125 mM KOAc, and protease inhibitor cocktail). 4. Resuspend cells in 1 volume of pellet/5 volumes of homogenization buffer. 5. Cells are then broken in a stainless ball-bearing homogenizer (clearance 18 µm) by passing the cell suspension through the homogenizer four or five times. 6. A postnuclear supernatant is created by centrifuging at 800 × g for 5 min at 4˚C. 7. This postnuclear supernatant is further centrifuged at 100,000 × g for 1 hr at 4˚C. 8. The supernatant is collected and passed through a Sephadex G-25 column pre-equilibrated in homogenization buffer to remove small molecules such as nucleotides. 9. Aliquots of 50 µl are frozen in liquid nitrogen after protein concentration determined by the Bradford assay and cytosol (~2 mg protein/ml) is stored at –80˚C for up to 6 months.

6.2.4 PREPARATION OF ENERGY-REGENERATING SYSTEM FOR CELL-FREE RECONSTITUTION SYSTEM An energy-regenerating system (ERS) consists of 1 mM ATP, 0.02 mM GTP, 12 mM creatine phosphate (CP), and 80 µg/ml creatine phosphokinase (CPK). 1. Aliquots of 100 mM ATP, 2 mM GTP, 600 mM CP, and 8 mg/ml CPK are prepared from powder and stored at –20˚C in water for later use. 2. To make 20X ERS, first mix ATP, GTP, and CP concentrated aliquots at a volume ratio of 1:1:2. 3. Titrate the pH of this mixture to 7 by adding a few µl of 1 N KOH and check the pH with pH paper. 4. Add one volume of CPK into the ERS mixture. 5. Add 15 µl of 20X ERS to a 300-µl cell-free reaction system.

β GENERATION IN CELL-FREE SYSTEM UTILIZING ERS 6.2.5 Aβ IN THE ABSENCE OF CYTOSOL 1. Permeabilized cells (cell-free system) are suspended in breaking buffer with a final volume of 300 µl containing 25 mM HEPES, 2.5 mM MgCl2, 0.5 µM CaCl2, and 110 mM KCl.

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2. Additionally, 15 µl of 20X ERS is added. The final concentration of ERS in the cell-free system will be 1 mM ATP, 0.02 mM GTP, 12 mM creatine phosphate, and 80 µg/ml creatine phosphokinase. 3. Vortex the reactions briefly to mix cells and reagents. 4. Incubate the cell-free reactions at 37˚C in a water bath for 90 min. 5. Gently vortex the reaction tubes every 15 min during incubation.

6.2.6 DETECTION

OF

β PRODUCTION Aβ

FROM

CELL-FREE SYSTEM

1. For Aβ detection in a cell-free system, spin reaction samples at 12,500 rpm for 30 sec after incubation period. 2. Transfer supernatant to new Eppendorf tubes and add 100 µl of 3% SDS (with 8 µl β-mercaptoethanol/ml SDS solution) to each pellet. For supernatant, add 15 µl of 10% SDS to make a final of 0.5% SDS. 3. Vortex at full speed at 95oC for 5 min in an Eppendorf Thermomixer. 4. Let samples cool down at room temperature. 5. Sonicate pellet samples to shear DNA or facilitate sample solubilization when necessary. 6. Add 1 ml immunoprecipitation (IP) buffer (50 mM Tris HCl, pH 8.8, 150 mM NaCl, 6 mM EDTA, 2.5% Triton X-100, 5 mM methionine and cysteine, and 1 mg/ml bovine serum albumin) into all fractions and vortex. (Alternatively, an IP buffer consisting of 20 mM Tris HCl, pH 7.4, 300 mM NaCl, 5 mM EDTA, and 1% Triton X-100 is equally effective.) 7. Spin at 12,000 × g or at full speed for 10 min at 4oC in a Brinkman centrifuge and collect supernatants. This step is absolutely necessary to eliminate an otherwise strong and ubiquitous background. 8. Immunoprecipitate Aβ using antibody 4G8 (Signet Laboratories, Inc., Dedham, MA). 9. Immunoprecipitated Aβ can be resolved by 10 to 20% tricine gels and then transferred to a 0.2 µm PVDF membrane and analyzed by autoradiography.

6.2.7 FORMATION OF NASCENT SECRETORY VESICLES IN CELL-FREE SYSTEM SUPPLEMENTED WITH ERS AND CYTOSOL 1. Permeabilized cells (cell-free system) will be incubated in a final volume of 300 µl containing 2.5 mM MgCl2, 0.5 µM CaCl2, and 110 mM KCl, an energy-regenerating system consisting of 1 mM ATP, 0.02 mM GTP, 10 mM creatine phosphate, 80 µg/ml creatine phosphokinase, and a protease inhibitor mixture. 2. 15 µl cytosol prepared from N2a cells16,19 at a concentration of 1 to 2 µg/µl (15 to 30 µg proteins) is included in the cell-free system to initiate nascent vesicle budding from the TGN membrane or ER membrane. 3. Incubations are carried out at 37oC to initiate nascent vesicle release. 4. Cell-free systems are incubated for various periods (15 to 120 min) to observe the kinetics of protein trafficking and the production of Aβ from both membrane and vesicle fractions.

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N

Hypotonic Treatment

N

N

Scraping

10 min labeling + 2 h incubating at 20° Or 4h labeling at 15°

Cytosol, Energy, Ions, etc.

Incubation at 37° or 20° N

ER markers (Calnexin, BiP) TGN markers (TGN38, γ Adaptin, Sialyl-Tase)

Sucrose Gradients Fractionation

N

Centrifugation Supernatant (Nascent Vesicles)

Pellet (Organelles)

β PP or A β detection (IP, SDS-PAGE, Mass-Spec.)

FIGURE 6.1 Cell-free reconstitution system. Cells are labeled with [35S]methionine for 10 min and then incubated at 20˚C, a temperature at which transport of proteins, including βAPP, from the ER to the TGN is unimpaired. However, under these conditions, the egress of secretory vesicles from the TGN is blocked, thus allowing labeled βAPP (and other membrane proteins) to accumulate in the TGN.3,5,16 This experimental design assures that TGN-specific vesicle biogenesis is measured after reconstitution of the cell-free system. Alternatively, cells are labeled with [35S]-methionine at 15˚C to accumulate βAPP within the ER. Cells are then permeabilized, followed by incubation at 37˚C to initiate vesicle release. Following incubation, vesicle and membrane fractions are separated by centrifugation at 11,000 rpm for 30 seconds at 4oC in a Brinkman centrifuge. Vesicle (supernatant) and membrane (pellet) fractions are diluted with IP buffer, and immunoprecipitated using antibody against βAPP or Aβ and analyzed by SDS-PAGE. ER and TGN membranes can be separated by sucrose gradient fractionation and verified by specific markers.

5. Following incubation of cell-free systems, vesicle and membrane fractions can be separated by centrifugation at 11,000 × g for 30 sec at 4oC in a Brinkman centrifuge. 6. The integrity of TGN stacks, ER membrane and derived vesicles can be demonstrated by electron microscopy (Figure 6.1).

6.2.8 DETECTION OF βAPP IN DIFFERENT FRACTIONS IMMUNOPRECIPITATION

THROUGH

1. Resuspend vesicle (supernatant) and membrane (pellet) fractions with 1 ml IP buffer (50 mM Tris HCl, pH 8.8, 150 mM NaCl, 6 mM EDTA, 2.5% Triton X-100, 5 mM methionine and cysteine, and 1 mg/ml bovine serum albumin). 2. Centrifuge resuspended fractions at 14,000 rpm for 10 min at 4oC in a Brinkman centrifuge.

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3. Collect supernatants and immunoprecipitate full-length βAPP and C-terminal fragments (CTFs) using antibody 3697,15 or antibody 4G8. 4. Immunoprecipitated βAPP can be resolved in 4 to 12% Tris–glycine gels and analyzed by autoradiography. CTFs can be resolved in 10 to 20% Tricine gels and analyzed by autoradiography.

6.3 DISCUSSION In summary, the procedures described herein provide detailed information about the preparation of a cell-free reconstitution system for studying intracellular Aβ production as well as trafficking of βAPP through secretory compartments. Figure 6.1 shows the outline of the experimental procedures. The discussion that follows focuses on specific areas in detail.

β GENERATION 6.3.1 ATP-DEPENDENT Aβ RECONSTITUTION SYSTEM

IN

CELL-FREE

Because our cell-free system allows us to reconstitute Aβ generation and produce various C-terminal fragments of APP, we studied both β-secretase and γ-secretase activities independently and dissected the requirements and optimal conditions for both. We showed that ATP is required for γ-secretase activity to yield Aβ (Figure 6.2). If the ATP-degrading enzyme apyrase was included in the cell-free system at 37oC, much less Aβ was generated compared to that in which an ATP regenerating system was present. In addition, we demonstrated that while ATP is required for γ-secretase activity, it appears less essential for β-secretase activity. Therefore, supplementation of an energy-regenerating system in the cell-free reconstitution assay is crucial to study γ-secretase-dependent Aβ generation.

a.

b.

1.0 β−CTF

βCTF(C99)

14.3

0.5

6.5

Aβ

3.4 20

°C

+A

py ra

se

+A TP

Aβ –0.5

19°

APY

ATP

FIGURE 6.2 Gamma-secretase cleavage of βAPP in the cell-free reconstitution system requires ATP. (a) Aβ generation is reconstituted in the TGN which is incubated at 37oC in the presence of an energy regenerating system (+ATP) or without an ATP regenerating system in the presence of the ATP degrading enzyme apyrase (+Apyrase). (b) Quantitative analysis of (a).

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6.3.2 CYTOSOL-DEPENDENT βAPP TRAFFICKING FROM TGN/ER IN THE CELL-FREE RECONSTITUTION SYSTEM The importance of cytosolic trafficking proteins in the genesis and budding of TGN vesicles is well established.26 It was not known, however, whether these trafficking factors are regulated by estrogen. Utilizing the cell-free trafficking assay, we demonstrated the effects of estrogen on cytosolic regulation of TGN vesicle biogenesis.16 Cytosol prepared from N2a cells stimulated TGN vesicle budding between 1 ng of protein/ml and 1 mg of protein/ml with maximal stimulation reached at 100 ng of protein/ml (Figure 6.3a). Cytosol prepared from cells treated with estrogen (estrogen-primed cytosol) stimulated budding nearly two times when compared with reactions using cytosol prepared from control cells (estrogen-naive cytosol) (Figure 6.3b, lane 5 vs. lane 7) and nearly two and a half times when compared to identical reactions without cytosol (Figure 6.3b, lane 3 vs. lane 7). These results suggest that supplementing cytosol in the cell-free reconstitution system is necessary to initiate vesicle release from the secretory compartments and that estrogen treatment stimulates protein transport from the TGN through alteration of cytosol/TGN membrane composition.

6.3.3 UTILIZING THE CELL-FREE RECONSTITUTION SYSTEM TO DEMONSTRATE PS1-REGULATED INTRACELLULAR TRAFFICKING AND SURFACE DELIVERY OF βAPP The formation of βAPP-containing vesicles from the TGN and from the ER in PS1-/fibroblasts was assessed, by comparison to wild-type (wt) cells, using the cell-free reconstitution system. As shown in Figure 6.4, budding of βAPP-containing vesicles from the TGN or from the ER was greatly increased at all time points examined in preparations that lacked PS1 when compared to preparations from cells that express wt PS1 (Figure 6.4a and Figure 6.4b). Moreover, live staining of loss-of-function PS1 cells (∆M1,2)29 using monoclonal antibody 6E10 revealed an obvious increase in the amount of surface-bound βAPP compared to PS1 wt cells (Figure 6.4c). The amounts of total surface glycoproteins are identical in the two types of cells as judged by staining for Vicia villosa agglutinin (VVA). The amount of newly synthesized βAPP delivered to the cell surface was further measured quantitatively in these cells by pulse-chase labeling in combination with cell surface biotinylation27 in intact cells. As shown in Figure 6.4d, up to 14.3% of nascent βAPP was transported to the plasma membrane after 120 min of chase, a value that is 48.9% greater than that in PS1 wt cells.

6.3.4 CHARACTERIZATION OF VESICLE AND MEMBRANE FRACTIONS BY SUCROSE GRADIENT AND ELECTRON MICROSCOPY Following permeabilization and incubation, the cell-free reconstitution system can be separated into vesicle and TGN/ER membrane fractions. The nascent vesicles as well as the intact TGN stacks and ER membranes can be determined by electron

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6

c.

5

+ 17β-E2

4 3

-17β-E2

2 1 0

βAPP Vesicle Budding (Arbitrary Units)

0

b.

βAPP Vesicle Budding (Arbitrary Units)

βAPP Vesicle Budding (Arbitrary Units)

a.

3.0 2.5

Control cytosol

5

Estrogen cytosol

4 3 2 1 0

10?g/ml 1ng/ml 100ng/ml 1mg/ml Cytosol protein concentration

Control cells 17β-E2 treated cells

6

Human 1 2

3

Rat 4

5

Mouse 6

Cytosol Source (cortical neurons)

d.

2.0

Rat Cortical Neurons

Mouse Cortical Neurons

20°C 37°C 37°C

20°C 37°C 37°C

1.5 1.0 17β-E2 treatment:

0.5

+

-

+

+

-

+

0 20°C 1 2

-Cyt. 3 4

17β-E2 Cyt. +Cyt. 5 6 7 8

Cytosol (100 ng/ml)

FIGURE 6.3 Cytosolic factors are up-regulated in response to 17β-E2. (a) Cell-free βAPP budding assays in the presence or absence of cytosol prepared from cells incubated with 17β-E2. Titration of cytosol demonstrates an increase in βAPP budding at all concentrations when using estrogen-primed cytosol versus estrogen-naive cytosol, with a maximum effect at 100 ng of protein/ml. (b) βAPP budding assays using no cytosol (–Cyt., lanes 3 and 4), estrogen-naive cytosol (lanes 5 and 6), and estrogen-primed cytosol (lanes 7 and 8) in untreated (odd lanes) or estrogen-treated (even lanes) cells. (c) Cytosol derived from primary human (lanes 1 and 2), rat (lanes 3 and 4), and mouse (lanes 5 and 6) neurons after incubation in either the absence (odd lanes) or presence (even lanes) of estrogen was used to stimulate βAPP budding in N2a cell-free assays. (d) Cell-free assays were performed in rat (left panel) and mouse (right panel) primary neurons. In each type, cells were incubated in either the absence or presence of 200 nM 17β-E2 for 1 wk before cell-free assays were performed. The budding of βAPP was assayed using 369 as described in methods. Experiments performed at 20˚C provided negative control.

microscopy (shown in Figure 6.1). Furthermore, ER and TGN membranes can be separated by sucrose gradients fractionation and verified by organelle-specific markers. In summary, the methods described in this chapter allow quantitative study of the kinetics of TGN- and ER-specific βAPP vesicle budding. By taking advantage of the fact that secretory pathway transit can be restricted at specific points by temperature blocks and then reinitiated by removing those blocks, we successfully demonstrated that budding of vesicles carrying βAPP as cargo diminishes the substrate pool from which Aβ peptides can be derived.16 Utilizing the cell-free reconstitution

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a.

b. TGN Membrane 0’

15’ 30’ 60’ 90’

TGN Vesicles

ER Membrane

0’ 15’ 30’ 60’ 90’

0’

PS1WT

PS1WT

PS1-/-

PS1-/-

6E10

c.

FITC-VVA

15’ 30’ 60’ 90’

ER Vesicles 0’ 15’ 30’ 60’ 90’

d. Total Cell βAPP

WT

Newly Synthesized βAPP on Cell Surface

WT

∆Μ1,2 ∆Μ1,2 Chase Time (min)

0’ 10’ 20’ 30’ 40’ 50’ 60’ 120’

FIGURE 6.4 (See color insert following page 114.) PS1 deficiency or loss of function accelerates βAPP trafficking from the TGN/ER and increases cell surface delivery. (a) and (b) Cell-free βAPP budding assays were prepared from PS1–/– fibroblasts expressing human βAPPswe alone (PS1–/–) or coexpressing βAPPswe and wild-type human PS1 cells. Cells were first labeled for 15 min with [35S]methionine at 37˚C and chased for 2 hr at 20˚C to accumulate labeled βAPP in the TGN. Alternatively, cells were labeled for 4 hr at 15˚C to accumulate labeled βAPP within the ER. Permeabilized cells were prepared and incubated at 37˚C for various periods to allow the formation of post-TGN or post-ER vesicles. (c) Live N2a cells were incubated with primary antibody 6E10 (1:100) at 4˚C for 1 hr to label cell surface βAPP (shown in red in color insert), and FITC-conjugated VVA to stain all surface glycoproteins (shown in green in color insert). Cells were then fixed and visualized by confocal microscopy. (d) Cells were labeled with [35S]methionine at 37oC for 10 min and chased at 20˚C for 2 hr to accumulate labeled βAPP in the TGN (total cell βAPP). Cells were then incubated at 37oC for various periods to allow transport of βAPP to the plasma membrane. Cell surface proteins were then biotinylated at 4˚C for 15 min. Biotinylated and nonbiotinylated proteins were separated into two fractions by binding to streptavidin beads. βAPP was immunoprecipitated from each fraction and analyzed.

system, the study of intracellular Aβ generation and regulation of βAPP trafficking through secretory compartments becomes feasible, as does analysis of other Alzheimer’s-associated secretory molecules.

ACKNOWLEDGMENTS This work was supported by grants from the National Institutes of Health (F32 AG23431 to DM and NS046673 to HX) and the Alzheimer’s Association (to HX).

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REFERENCES 1. Selkoe, D.J. The cell biology of beta-amyloid precursor protein and presenilin in Alzheimer’s disease. Trends Cell Biol. 8, 447–453, 1998. 2. Greenfield, J.P. et al. Cellular and molecular basis of beta-amyloid precursor protein metabolism. Front. Biosci. 5, D72–D83, 2000. 3. Xu, H. et al. Generation of Alzheimer beta-amyloid protein in the trans-Golgi network in the apparent absence of vesicle formation. Proc. Natl. Acad. Sci. USA 94, 3748–3752, 1997. 4. Hartmann, T. et al. Distinct sites of intracellular production for Alzheimer’s disease A beta40/42 amyloid peptides. Nat. Med. 3, 1016–1020, 1997. 5. Greenfield, J.P. et al. Endoplasmic reticulum and trans-Golgi network generate distinct populations of Alzheimer beta-amyloid peptides. Proc. Natl. Acad. Sci. USA 96, 742–747, 1999. 6. Sisodia, S.S. Beta-amyloid precursor protein cleavage by a membrane-bound protease. Proc. Natl. Acad. Sci. USA 89, 6075–6079, 1992. 7. Caporaso, G.L. et al. Morphologic and biochemical analysis of the intracellular trafficking of the Alzheimer beta/A4 amyloid precursor protein. J. Neurosci. 14, 3122–3138, 1994. 8. Nordstedt, C. et al. Identification of the Alzheimer beta/A4 amyloid precursor protein in clathrin-coated vesicles purified from PC12 cells. J. Biol. Chem. 268, 608–612, 1993. 9. Soriano, S. et al. Expression of beta-amyloid precursor protein-CD3gamma chimeras to demonstrate the selective generation of amyloid beta(1-40) and amyloid beta(1-42) peptides within secretory and endocytic compartments. J. Biol. Chem. 274, 32295–32300, 1999. 10. Perez, R.G. et al. Mutagenesis identifies new signals for beta-amyloid precursor protein endocytosis, turnover, and the generation of secreted fragments, including A beta42. J. Biol. Chem. 274, 18851–18856, 1999. 11. Cook, D.G. et al. Alzheimer’s A beta(1-42) is generated in the endoplasmic reticulum/intermediate compartment of NT2N cells. Nat. Med. 3, 1021–1023, 1997. 12. Buxbaum, J.D. et al. Processing of Alzheimer beta/A4 amyloid precursor protein: modulation by agents that regulate protein phosphorylation. Proc. Natl. Acad. Sci. USA 87, 6003–6006, 1990. 13. Xu, H. et al. Metabolism of Alzheimer beta-amyloid precursor protein: regulation by protein kinase A in intact cells and in a cell-free system. Proc. Natl. Acad. Sci. USA 93, 4081–4084, 1996. 14. Zheng, H. et al. Modulation of A (beta) peptides by estrogen in mouse models. J. Neurochem. 80, 191–196, 2002. 15. Xu, H. et al. Estrogen reduces neuronal generation of Alzheimer beta-amyloid peptides. Nat. Med. 4, 447–451, 1998. 16. Greenfield, J.P. et al. Estrogen lowers Alzheimer beta-amyloid generation by stimulating trans-Golgi network vesicle biogenesis. J. Biol. Chem. 277, 12128–12136, 2002. 17. Xu, H. and Shields, D. Prohormone processing in the trans-Golgi network: endoproteolytic cleavage of prosomatostatin and formation of nascent secretory vesicles in permeabilized cells. J. Cell Biol. 122, 1169–1184, 1993. 18. Ling, W.L., Siddhanta, A., and Shields, D. The use of permeabilized cells to investigate secretory granule biogenesis. Methods 16, 141–149, 1998.

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19. Musch, A. et al. Transport of vesicular stomatitis virus G protein to the cell surface is signal mediated in polarized and nonpolarized cells. J. Cell. Biol. 133, 543–558, 1996. 20. Rothman, J.E. Mechanisms of intracellular protein transport. Nature 372, 55–63, 1994. 21. Sweeney, D.A., Siddhanta, A., and Shields, D. Fragmentation and re-assembly of the Golgi apparatus in vitro. A requirement for phosphatidic acid and phosphatidylinositol 4,5-bisphosphate synthesis. J. Biol. Chem. 277, 3030–3039, 2002. 22. de Silva, A.M., Balch, W.E., and Helenius, A. Quality control in the endoplasmic reticulum: folding and misfolding of vesicular stomatitis virus G protein in cells and in vitro. J. Cell Biol. 111, 857–866, 1990. 23. Doms, R.W. et al. Role for adenosine triphosphate in regulating the assembly and transport of vesicular stomatitis virus G protein trimers. J. Cell. Biol. 105, 1957–1969, 1987. 24. Netzer, W.J. et al. Gleevec inhibits beta-amyloid production but not Notch cleavage. Proc. Natl. Acad. Sci. USA 100, 12444–12449, 2003. 25. Beckers, C.J. and Balch, W.E. Calcium and GTP: essential components in vesicular trafficking between the endoplasmic reticulum and Golgi apparatus. J. Cell Biol. 108, 1245–1256, 1989. 26. Sollner, T.H. and Rothman, J.E. Molecular machinery mediating vesicle budding, docking and fusion. Experientia 52, 1021–1025, 1996. 27. Yan, R. et al. The transmembrane domain of the Alzheimer’s beta-secretase (BACE1) determines its late Golgi localization and access to beta-amyloid precursor protein (APP) substrate. J. Biol. Chem. 276, 36788–36796, 2001. 28. Xu, H., Greengard, P., and Gandy, S. Regulated formation of Golgi secretory vesicles containing Alzheimer beta-amyloid precursor protein. J. Biol. Chem. 270, 23243–23245, 1995. 29. Leem, J.Y. et al. A role for presenilin 1 in regulating the delivery of amyloid precursor protein to the cell surface. Neurobiol. Dis. 11, 64–82, 2002.

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7

Studying Amyloid β-Protein Assembly Erica A. Fradinger, Samir Kumar Maji, Noel D. Lazo, and David B. Teplow

CONTENTS 7.1 7.2 7.3

7.4

Introduction Background Peptide Production and Preparation 7.3.1 Production of Aβ Peptides 7.3.2 Preparation of Starting Peptide Stocks 7.3.2.1 HFIP Pretreatment 7.3.2.2 NaOH Pretreatment 7.3.2.3 Sample Clarification and Fibril Isolation 7.3.2.4 Preparation of LMW Aβ by Size Exclusion Chromatography 7.3.2.5 Preparation of LMW Aβ by Filtration 7.3.2.6 Preparation of Aggregate-Free Aβ by Ultracentrifugation 7.3.2.7 Preparation of Oligomeric Aβ42 7.3.2.8 Preparation of Aβ Protofibrils 7.3.2.9 Preparation of Aβ Fibrils Monitoring Aβ Assembly. 7.4.1 Oligomerization 7.4.1.1 Determination of Oligomer Size Distributions Using PICUP 7.4.1.2 Determination of Oligomer Size Using SEC 7.4.1.3 Determination of Particle Diffusion Coefficients Using QLS 7.4.2 Fibril Formation 7.4.2.1 Thioflavin T (ThT) Binding 7.4.2.2 Congo Red Binding 7.4.2.3 Turbidity 7.4.3 Secondary Structure Determination 7.4.3.1 CD Spectroscopy 7.4.3.2 Fourier Transform Infrared Spectroscopy (FTIR)

0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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7.4.4

Topographical Analysis 7.4.4.1 8-Anilino-1-Naphthalenesulfonic Acid (ANS) Binding 7.4.4.2 Intrinsic Fluorescence 7.4.4.3 Electron Paramagnetic Resonance (EPR) 7.4.4.4 Hydrogen–Deuterium Exchange 7.4.5 NMR Spectroscopy 7.4.6 Morphological Analysis 7.4.6.1 Electron Microscopy 7.4.6.2 Atomic Force Microscopy 7.5 Discussion Acknowledgments References

7.1

INTRODUCTION

The amyloid β-protein precursor (AβPP), also referred to as the amyloid precursor protein (APP), is a type I transmembrane glycoprotein.1 Through the actions of specific endoproteases, the ~110 to 135 kDa AβPP is post-translationally processed, producing a 40- to 42-amino acid amphipathic peptide, the amyloid β-protein (Aβ).2 Aβ is expressed ubiquitously in the human body as a normal part of cellular and organismal physiology.3,4 Pioneering work by George Glenner two decades ago5,6 revealed that the protein deposits that are pathognomonic for Alzheimer’s disease (AD) are composed primarily of Aβ. Since then, tremendous advances have been made in our understanding of both Aβ and AβPP. Genetic evidence revealed a causal link between AβPP and AD.1 In all kindreds thus far examined, mutations in genes encoding AβPP or in genes encoding proteins involved in AβPP metabolism either cause increased production of the longer, more amyloidogenic, 42-residue form of Aβ (Aβ42) or production of Aβ peptides containing amino acid substitutions affecting Aβ self-assembly.7 Aβ assembly thus is inextricably linked with AD. For this reason, therapeutic strategies for AD have focused on controlling Aβ metabolism, Aβ-mediated neurotoxicity, or Aβ self-assembly.8 The centrality of Aβ in the neuropathogenesis of AD has made biochemical and biophysical studies of Aβ aggregation exceedingly important. How is this done? The goal of this chapter is to provide readers the answer to this question, specifically with respect to studies of Aβ folding and self-assembly. Moreover, Aβ assembly is an archetype for amyloid assembly. Therefore, the methods discussed here are relevant and applicable to the study of many other amyloidogenic proteins and peptides, of which ~25 have been defined clinically.

7.2 BACKGROUND The assembly of Aβ from a nascent monomer into the structure known as amyloid 9–11 is an exceedingly complex process.12–14 In addition to the seminal event of Aβ selfassociation, other proteins, glycosaminoglycans, lipids, and reactive oxygen species also may affect amyloidogenesis. Ideally, one would like to construct an assembly scheme incorporating all elements that contribute significantly to the process. To do

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so, reductionist approaches seek to understand each element with great mechanistic detail. Logically, the most important element is Aβ itself. Determining the tertiary structures of Aβ in its monomeric and assembled states, the quaternary structures resulting from monomer self-association and the dynamics of the association processes are important goals. These data must be correlated with the biological behaviors of specific assemblies in order to determine meritorious therapeutic targets. A prerequisite for any study of Aβ assembly is pre facto consideration of which assembly is to be studied. Aβ forms a variety of structures, including multiple monomeric conformers,15 different types of oligomers,16–18 Aβ-derived diffusible ligands (ADDLs),19,20 annuli,21 micelles,22,23 protofibrils,24,25 fibrils,12 and spheroids.26,27 The structural relationships among these assemblies and the differences in the assembly processes of wild type Aβ40, Aβ42, and mutant Aβ peptides are not entirely understood and are areas of active investigation. Studies of Aβ comprise three different classes — static, dynamic, and biological. Static studies reveal features of the assembly process observable at defined times, e.g., the morphology of end-stage structures such as fibrils. Electron microscopy (EM), x-ray diffraction, solid state nuclear magnetic resonance spectroscopy (NMR), and some types of atomic force microscopy (AFM) are examples of useful techniques for studying static features of Aβ assembly. Dynamic studies reveal time-dependent changes in peptide conformation and aggregation state. Dynamics studies can be accomplished through continuous monitoring or iterative use of static techniques. Circular dichroism (CD) spectroscopy, Congo Red and thioflavin T dye binding, turbidity, quasielastic light scattering spectroscopy (QLS), hydrogen–deuterium exchange, electron paramagnetic resonance spectroscopy (EPR), NMR, size exclusion chromatography (SEC), intrinsic fluorescence, Fourier transform infrared spectroscopy (FTIR), and analytical ultracentrifugation are examples of techniques for studying assembly dynamics. Biological studies elucidate assembly-specific effects on cellular function and typically involve vital assays (cell death) and metabolic assays (redox activity, transcriptional dynamics, signaling events, and post-translational modifications). The two most important questions common to all of these approaches are (1) how does one initially prepare the peptide for study and (2) how does one perform each assay? The importance of question 1 with respect to the proper interpretation of data produced subsequently cannot be overemphasized. It also should be noted that implicit in question 2 is the issue of how structure–activity relationships can be defined in studies of heterogeneous structures existing in rapid equilibria. In the sections that follow, we discuss issues associated with preparation of peptide stocks and provide selected examples of static and dynamic approaches for monitoring early, mid-, and late-stage events in Aβ assembly.

7.3 PEPTIDE PRODUCTION AND PREPARATION 7.3.1 PRODUCTION

OF

Aβ PEPTIDES

Aβ is produced either by solid-phase peptide synthesis (SPPS) or through recombinant DNA techniques, providing high purity peptide suitable for in vitro and in vivo

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experimentation. Unfortunately, substantial compositional variation has been reported among Aβ preparations, resulting in experimental irreproducibility.28–31 It is important that the experimentalist verifies that the peptide itself is chemically pure and that nonpeptide components are either absent from peptide stocks or inert. Most peptide lyophilizates are not 100% peptide by weight, but contain salts and other components. For all the protocols that follow, when calculating initial peptide concentration, peptide weight (and not total lyophilizate weight) must be used. Intrinsic chemical purity is the state in which the primary structure (protein sequence) of Aβ is correct. SPPS cannot produce an Aβ product that is 100% pure. Failure sequences, peptides missing one or more amino acids, are unavoidable, although with proper synthesis chemistry their relative amounts can be minimized. Oxidation of Met35 to its corresponding sulfoxide, Met(O)35, is a common reaction that can occur during peptide workup and purification, especially in the presence of formic acid or oxidants. Synthesis-related amino acid racemization, and side-reactions during peptide cleavage and deprotection, may also be observed, but these generally occur infrequently. Most peptide suppliers perform quantitative amino acid analysis and mass spectrometry to characterize their products. However, because amino acid and simple mass analyses cannot determine primary structure, Edman or mass spectrometric sequence analysis can be used to prove formally that the peptide structure is correct. With respect to nonpeptide components of peptide preparations, fluoren-9-ylmethoxycarbonyl (FMOC)-mediated SPPS and high performance liquid chromatography (HPLC) purification produce trifluoroacetic acid (TFA) salts of the resulting peptides. These salts along with chemical scavengers are often present in lyophilized peptide preparations and can complicate the initial solvation and preparation of peptide stock solutions. For recombinantly derived Aβ, primary structure changes are rare because of the high fidelity of the protein expression systems and the physiologic conditions under which these systems operate. In these systems, Aβ often is produced as a fusion protein requiring post-translational processing with highly specific endoproteases. It is important to ensure that the Aβ component of the fusion protein is not contaminated by uncleaved fusion protein, the enzyme itself or by peptide fragments produced through adventitious proteolysis. Because fusion protein cleavage is performed with biological buffers, buffer exchange or removal may be necessary prior to peptide use.

7.3.2 PREPARATION

OF

STARTING PEPTIDE STOCKS

One of the most common problems encountered in work with Aβ is irreproducible behavior of the peptide. Primary causes of irreproducibility are the initial structure and aggregation state of the peptide, both in the solid state31 and immediately after solvation. Several solvation methods have been developed in an effort to eliminate preexisting aggregates and create conformationally uniform, monomeric, Aβ stock solutions. A “magic formula” for achieving this goal has not been found. Dimethyl sulfoxide (DMSO),32–34 TFA,35 trifluoroethanol (TFE),36 hexafluoroisopropanol (HFIP),33,37 and NaOH31 all have been used to solubilize Aβ. HFIP and TFE disrupt hydrophobic interactions in aggregated amyloid preparations and stabilize α-helical

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structure,36,38,39 leading to disruption of preexistent β-sheet structure. HFIP pretreatment (Section 7.3.2.1) of Aβ has been shown to yield peptide solutions of uniform globular morphology with predominantly α-helical and random coil secondary structures and less than 1% β-sheet.33 Aβ prepared by SPPS can contain residual TFA. Dissolution of these peptide lyophilizates in water produces a strongly acidic solution (pH 3 to 4). Addition of biological buffers of neutral pH causes the solution pH to pass through the isoelectric point of Aβ (~5.5) — a point at which the solubility of Aβ is minimal and its propensity to aggregate is maximal.38 The result is rapid aggregation that produces an inhomogeneous starting peptide solution. One strategy that addresses this problem is to predissolve Aβ under strongly alkaline conditions and then to relyophilize the solutions prior to solvation in biological buffers. NaOH pretreatment (Section 7.3.2.2) has been shown to improve significantly the yield of unaggregated Aβ.31 A number of postsolvation approaches have been developed to remove preexisting aggregates. Filtration methods have been used to remove large and small aggregates from starting solutions of Aβ.31 Filtration of Aβ through a 0.2-µm nylon Micro-Spin Whatman filter at 5000 × g for 10 min will remove fibrils, fibril aggregates, and other structures larger than 200 nm. However, for most experimental needs, this filtration is insufficient because assemblies of 200-nm size are quite large and generally are already fibrillar. Filtration through filters with 10-kDa exclusion limits (see Section 7.3.2.5) is a superior method that initially yields monomers and dimers. It is important to note that, in peptide regimes of µM concentration and higher, nascent Aβ monomers immediately establish a rapid equilibrium with higher order oligomers.17,40 Most aggregate-free Aβ solutions therefore comprise an oligodisperse population of assemblies. Nevertheless, because this population can be produced routinely and is not polydisperse, reproducible peptide assembly behavior can be observed. Another highly effective procedure for preparing aggregate-free Aβ is SEC (Section 7.3.2.4). SEC allows the almost simultaneous collection of larger oligomeric populations, including protofibrils (Section 7.3.2.8).24 Although more laborious, ultracentrifugation (Section 7.3.2.6) also has been used to prepare aggregate-free, nominally monomeric, peptide stock solutions.41 In the sections that follow, we present protocols for solvating Aβ lyophilizates, clarifying turbid solutions (Section 7.3.2.3), producing aggregate-free Aβ solutions, isolating oligomeric assemblies (Section 7.3.2.7) and protofibrils, and preparing fibrils (Section 7.3.2.9). It should be noted that some controversy surrounds the oligomerization state of aggregate-free starting preparations of Aβ. Here we refer to these preparations, which typically exist at concentrations of 10 to 50 µM, as low molecular weight (LMW) Aβ.42 We do so because techniques including QLS,24 chemical cross-linking,17,40,43 fluorescence resonance energy transfer,44 and ultracentrifugation45,46 have shown that a monodisperse monomer population does not exist in these solutions. Readers should note that for many experimental needs this issue may be academic because the primary concern is the ability to prepare peptide stock solutions identical in their distribution of peptide assembly states, whatever that distribution may be.

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7.3.2.1 HFIP Pretreatment 1. 2. 3. 4.

Dissolve Aβ peptide in 100% HFIP to a final concentration of 0.5 mM. Distribute 100 µl aliquots into 1.5 ml polypropylene microcentrifuge tubes. Remove HFIP by evaporation under a gentle stream of nitrogen or argon. Remove residual HFIP in vacuo, using a lyophilizer or a centrifugal concentrator (e.g., Savant SpeedVac concentrator, Savant Instruments, Holbrook, NY). 5. The final product will be a peptide film within the microcentrifuge tube. If properly desiccated, the tubes can be stored for extended periods (months to years) at –20 or –80ºC.

Note: All work with HFIP should be conducted in a chemical fume hood with adequate protection. The peptide film produced by this procedure may not easily dissolve in water or aqueous buffers. Initial solvation in DMSO can facilitate complete resuspension. 7.3.2.2 NaOH Pretreatment 1. Suspend lyophilized Aβ in 2 mM NaOH at a concentration of 1 mg/ml. It is important to ensure that the pH of the resulting solution reaches at least 10.5.31 2. Sonicate the suspension in a bath-type sonicator for 1 min (#1200-R, Branson Ultrasonics, Danbury, CT). 3. Lyophilize the resulting NaOH-treated peptide. 4. Store the lyophilizate at –20°C until needed. 7.3.2.3 Sample Clarification and Fibril Isolation For some experiments, the investigator may want to fractionate or remove large aggregates from more buoyant structures, such as protofibrils, oligomers and LMW Aβ. A simple method is low speed centrifugation. Elimination of large, experimentally irrelevant or undesired aggregates produces a “clarified” supernatant fluid containing protofibrils and smaller assemblies. The pellet contains aggregates and fibrils that may be used for morphological studies as well as seeding experiments. 1. Dissolve Aβ at a concentration of 2 mg/ml in Milli-Q water and then add an equal volume of double-strength (2X) buffer. 2. Sonicate the sample for 1 min in an ultrasonic water bath (#1200-R, Branson). 3. Centrifuge the sample for 10 min at 16,000 × g using a bench top microcentrifuge. β by Size Exclusion Chromatography 7.3.2.4 Preparation of LMW Aβ 1. Prepare 10 mM sodium phosphate, pH 7.4, and then filter the solution through a 0.22-µm polyethersulfone (or equivalent) membrane to remove bacteria and any other large particulates.

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2. Using the buffer prepared in step 1, wash and equilibrate a 10/30 Superdex 75 HR (Amersham Biosciences, Piscataway, NJ) column at a flow rate of 0.5 ml/min until a flat ultraviolet (UV) trace is observed. The chromatographic pumping system per se is not relevant to the procedure. It need only provide appropriate flow rates and include a detector capable of determining absorbance in the UV range (200 to 300 nm). 3. Dissolve 350 to 400 µg of lyophilized Aβ in DMSO at a concentration of 2 mg/ml. 4. Sonicate the dissolved peptide for 1 min in a bath sonicator (#1200-R, Branson). 5. Centrifuge the peptide solution at 16,000 × g for 10 min to remove any large aggregates. 6. Inject 160 to 180 µl of the supernate onto the equilibrated column. 7. UV monitoring is used to detect peaks eluting from the column. We routinely monitor at 254 nm, but wavelengths of 215 nm (peptide bond absorbance) or 280 nm (tyrosine absorbance) are also satisfactory. A void volume peak elutes from the column first. A large peak composed of LMW Aβ elutes late in the run with a retention time consistent with globular standards of molecular mass 5 to 15 kDa. Column calibration both with globular and polymeric standards will provide the most accurate indications of apparent molecular weight. 8. Generally, only the apex (middle third, based on collection time) of the LMW peak is collected and used. Note: LMW Aβ comprises an equilibrium mixture of low-order oligomers. It is critical that this material be used immediately after its isolation if structure–activity correlations are required. Time delays and sample manipulation allow assembly of larger structures, which can complicate interpretation of the experimental data. β by Filtration 7.3.2.5 Preparation of LMW Aβ An alternative to SEC for preparation of LMW Aβ is filtration. This method is simpler than SEC and requires less time and fewer instrumental resources. The LMW Aβ40 produced is qualitatively similar to that produced by SEC. However, the oligomerization states of Aβ42 differ within LMW fractions prepared by the two methods.17 SEC-isolated LMW Aβ42 produces higher order oligomers, in addition to the relatively narrow (predominantly monomer through heptamer) distribution of oligomers observed with filtered preparations.17 1. Wash Centricon YM-10 filters with 200 µl of 10 mM phosphate buffer, pH 7.4, by adding the solution to the filter and centrifuging at 16,000 × g for 20 min. 2. Discard the filtrate and repeat step 1. 3. Dissolve lyophilized NaOH-treated (see section 7.3.2.2) Aβ in Milli-Q water at a concentration of 4 mg/ml. 4. Add an equal volume of 20 mM phosphate buffer, pH 7.4.

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5. Sonicate for 1 min (#1200-R, Branson). 6. Place the washed filter unit in a clean microcentrifuge tube, transfer the peptide solution to the unit, and then centrifuge at 16,000 × g for 30 min. 7. Collect the LMW Aβ filtrate. Note: The use of filtered deionized, 18.2-MΩ water is suggested. A Millipore Milli-Q Synthesis system is an excellent source for water. If the Aβ peptide available has not been NaOH-pretreated, pretreatment can be done prior to step 4 by adding 1 M NaOH to the peptide solution to produce a final NaOH concentration of 6 mM. For experiments involving extended incubation times, 0.01% (w/v) sodium azide is added to the buffers to prevent microbial growth. β by Ultracentrifugation 7.3.2.6 Preparation of Aggregate-Free Aβ Döbeli et al.41 developed a method for the production of “aggregation competent monomeric Aβ peptides.” This method uses fusion proteins produced in Escherichia coli and processed through chemical cleavage of the Aβ component, reduction of oxidized Met35, and multiple chromatographic steps. The final step, ultracentrifugation, is done immediately prior to use of the peptide to maximize the content of peptide monomer. This centrifugation produces soluble Aβ,46 equivalent in principle to LMW Aβ.42 Full details of the preparative and analytical ultracentrifugation procedures have been published.41,46 A brief protocol follows. 1. The starting peptide stock contains Aβ at a concentration of 50 to 100 µM in 12 mM Tris HCl, pH 8. The procedure is performed with Aβ42, but is applicable to any Aβ peptide. The buffer composition is not critical, as long as it does not accelerate Aβ self-association. Low ionic strength buffers without added salt (NaCl) are preferred. 2. The sample is introduced into the ultracentrifuge and spun at 320,000 × g for 12 hr at 20°C. 3. The supernate contains soluble Aβ estimated to contain >80% monomers. The remainder of the peptide mass comprises dimers and higher order oligomers. β42 7.3.2.7 Preparation of Oligomeric Aβ Aβ42 has been found to oligomerize in a distinctly different manner from Aβ40.17 In particular, Aβ42 has a propensity to form metastable globular structures approximately 5 to 6 nm in diameter. These have been termed paranuclei17 or ADDLs (Aβderived diffusible ligands).20 Aβ42 oligomers form immediately upon solubilization of synthetic Aβ42 preparations, but because these oligomers exist in a rapid equilibrium with monomers and larger structures, preparing well-defined oligomer populations is problematic. The following method details how to prepare ADDLs32 and is essentially identical to that used by Stein et al.33 As with all methods for preparing oligomeric or polymeric assemblies, the actual morphologies of the peptides should be confirmed by EM or AFM.

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1. 2. 3. 4. 5. 6. 7. 8.

Pretreat Aβ peptide with HFIP (Section 7.3.2.1). Resuspend peptide film in DMSO to a final concentration of 5 mM. Mix well by pipetting, then bath sonicate for 10 min (#1200-R, Branson). Add ice-cold phenol red-free F12 medium to the solution so that the final peptide concentration is 100 µM. Mix well by vortexing for 30 sec. Incubate at 4°C for 24 hr. Centrifuge at 14,000 × g for 10 min. The supernate contains ADDLs.

β Protofibrils 7.3.2.8 Preparation of Aβ 1. 2. 3. 4. 5.

Dissolve Aβ in Milli-Q water at a concentration of 4 mg/ml. Add an equal volume of 0.2 M Tris-HCl, pH 7.4. Incubate at room temperature (22°C) for 48 hr. Centrifuge the solution at 16,000 × g for 5 min. Remove the supernate and chromatograph on a Superdex 75 column, as described in Section 7.3.2.4. 6. The peak eluting immediately after the void volume of the column will contain protofibrils.

Note: The incubation time can be adjusted to provide the greatest yield of protofibrils. A 48-hr period generally produces equivalent amounts of protofibrils and LMW Aβ40, but times vary for Aβ42 and peptides containing amino acid substitutions associated with familial forms of AD. The method described here is the classic method for protofibril preparation. Solvent additives,47 alterations in buffer composition,48 and changes in the primary structure of Aβ49,50 also have been used to increase protofibril yield or stabilize the assembly. β Fibrils 7.3.2.9 Preparation of Aβ Fibril preparation is the easiest task to accomplish in studies of Aβ because fibril formation is the default pathway for Aβ assembly. Simple incubation of solubilized Aβ peptide at sufficient concentration in physiologic buffers will produce fibrils unless extraordinary precautions are taken to eliminate preexisting nidi for fibril nucleation. It should be noted, however, that many different polymeric structures have been observed, including short, stubby fibers, single filaments, twisted assemblies containing two or more subfilaments, ribbons, and sheets.51,52 Alterations in temperature, ionic strength, pH, solvent composition, and agitation can produce dramatic differences in the morphology of the end-stage assembly. Therefore, the final product should be examined by EM or AFM to determine its morphology. The protocol that follows is one means for obtaining fibrils. 1. Dissolve lyophilized Aβ in sterile Milli-Q water at a concentration of 4 mg/ml. 2. Add sufficient 1 M NaOH to produce a final concentration of 6 mM.

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3. Add an equal volume of 2× buffer. The buffer should be chosen based on the goal of the specific experiment conducted. 4. Sonicate the peptide sample for 10 min at room temperature (#1200-R, Branson). 5. Pellet any large aggregates at 16,000 × g for 10 min. 6. Transfer the supernate to a 1.5-ml microcentrifuge tube. 7. Incubate at 37°C. Agitation will accelerate the fibril formation process significantly.

β ASSEMBLY 7.4 MONITORING Aβ 7.4.1 OLIGOMERIZATION Data obtained from in vitro and in vivo studies correlating Aβ aggregation state and biological activity support the hypothesis that small, oligomeric Aβ assemblies may be key pathogenetic effectors of AD16,53–56 Therefore, understanding early Aβ assembly events is vital if rational approaches for AD therapy are to be developed. Achieving this understanding is complicated by the fact that assembly intermediates are metastable. Techniques such as EM and AFM are not ideal for elucidating assembly dynamics. The polydispersity of Aβ populations and the presence of large aggregates make QLS analyses difficult. SDS gel electrophoresis destroys noncovalent complexes, making this approach unsuitable. To allow study of assembly intermediates, methods have been sought for their stabilization. Recently, we introduced a method designated photo-induced crosslinking of unmodified proteins (PICUP) to the study of Aβ oligomerization.40,57 PICUP produces covalent bonds between unmodified peptides or proteins. To do so, a tris(bipyridyl)ruthenium(II) complex [Ru(Bpy)] is photo-oxidized in the presence of an electron acceptor. This produces Ru(III), a powerful one-electron oxidizer that abstracts electrons from reactive amino acids and results in free radical formation. The free radicals react quickly with neighboring groups, yielding covalently crosslinked peptides or proteins. PICUP has several attractive characteristics. It requires very short reaction times (1 sec) and no pre facto peptide modifications. It can be performed at neutral pH and under isotonic conditions. It is highly efficient (80%), and is initiated by visible (not UV) light, thereby preventing UV-induced peptide damage. We discuss here the use of PICUP to study peptide and protein oligomerization. 7.4.1.1 Determination of Oligomer Size Distributions Using PICUP 1. Prepare a 1 mM solution of tris(2,2′-bipyridyl)dichlororuthenium(II) [Ru(bpy)] in 10 mM sodium phosphate, pH 7.4. Protect this solution from ambient light. 2. Prepare 20 mM ammonium persulfate (APS) in 10 mM sodium phosphate, pH 7.4. 3. Prepare LMW Aβ. This can be done using SEC (Section 7.3.2.4), filtration (Section 7.3.2.5), or ultracentrifugation (Section 7.3.2.6).

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4. Add 18 µl of LMW Aβ into a thin-walled 0.2-ml polymerase chain reaction (PCR) tube. 5. Add 1 µl of Ru(bpy) and 1 µl of APS and mix well. 6. Illuminate for 1 sec. 7. Quench immediately by adding 1 µl of 1 M dithiothreitol in water. Note: We find that a convenient and inexpensive illumination system is a 35-mm SLR camera body to which a small bellows is attached. A 200-W incandescent lamp is an appropriate light source. This arrangement allows reproducible and precise illumination times. The sample is placed in the bellows, the bellows is capped, and the light source illuminates the shutter through the open film compartment (i.e., from the rear of the camera). Actuation of the shutter then allows the light to enter the bellows. Note also that instead of quenching with dithiothreitol, 5% (v/v) β-mercaptoethanol in SDS-PAGE sample buffer may be used. 7.4.1.2 Determination of Oligomer Size Using SEC SEC or gel permeation chromatography is a technique that fractionates solutes on the basis of their Stokes radii.58,59 SEC can be performed either in the presence or absence of denaturants. The absence of denaturants is advantageous relative to SDS gel electrophoresis because many oligomeric assemblies are SDS-labile. SEC is an excellent technique both for preparative work and for the analysis of particle size. The dynamic range of SEC is large, ranging from molecular weights below a thousand up to millions. However, the resolution is low, thus baseline separation of similarly sized proteins generally requires at least a 2× molecular weight difference. Nevertheless, SEC provides the means to isolate homogeneous fractions of particular Aβ assemblies and to monitor early Aβ oligomerization events. The basic instrumental arrangement and sample manipulations involved in SEC studies of Aβ have been described in Section 7.3.2.4. Iterative chromatographic analyses of Aβ assembly reactions can provide useful information on both the types of oligomers formed and the kinetics of oligomer formation and maturation into large polymers. An extensive literature exists in these areas, to which readers are referred to learn more about applications of SEC of special relevance to their experimental needs.17,24,27,40,50,60–62 7.4.1.3 Determination of Particle Diffusion Coefficients Using QLS QLS, also known as dynamic light scattering or photon correlation spectroscopy, is an optical method for the determination of diffusion coefficients of particles undergoing Brownian motion in solution.63,64 Diffusion coefficients are determined by particle size, shape, and flexibility as well as by interparticle interactions. These parameters provide important information about the kinetics and structural transitions in a system and can be studied by QLS. Using the Stokes-Einstein equation,65 D = kBT/6π ηRH , diffusion coefficients (D) can be converted into hydrodynamic radii (RH) — conceptually similar to Stokes radii. Thus QLS, like SEC, allows monitoring of protein assembly reactions. However, QLS provides a number of advantages. It is noninvasive; therefore reactions can be

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monitored in situ without the need for sample manipulation or consumption. QLS is rapid, requiring only minutes to acquire data, and is quantitative. QLS thus is an excellent tool for studying protein aggregation and has been used by a number of laboratories to examine Aβ assembly.17,22,61,66–69 In our laboratory, QLS studies have provided insight into nucleation and elongation rates of Aβ fibril formation,22,70 the existence and structural organization of micelle-like Aβ intermediates,22,71 activation energies for Aβ monomer addition to fibril ends,72 and sizes of the earliest Aβ42 oligomers (paranuclei).17 Although simple in principle, QLS requires substantial expertise in optical and polymer physics. A complete discussion of the subject is beyond the scope of this chapter and therefore interested readers are referred to a recent review on the theory and practice of QLS for studying protein aggregation.73

7.4.2 FIBRIL FORMATION 7.4.2.1 Thioflavin T (ThT) Binding ThT is a fluorescent dye that binds to β-sheet-rich assemblies.74,75 Excitation of the bound dye at 450 nm produces strong fluorescence at 482 nm, providing a means of monitoring formation of the extended sheets that comprise amyloid fibrils.76,77 1. Prepare a 100 µM stock solution of ThT in 10 mM phosphate buffer, pH 7.5. 2. Mix 5 µl of the ThT stock with 100 µl of an Aβ sample. 3. Record the fluorescence intensity four times, at 10-sec intervals after 90 sec of incubation. Use an excitation wavelength of 450 nm (slit width = 5 nm) and an emission wavelength of 482 nm (slit width = 10 nm). Average the four readings to obtain a value for the ThT fluorescence. 4. Determine the “blank-corrected” fluorescence by measuring the fluorescence of a peptide-free sample and subtracting this intensity from those of the peptide samples. Note: Iterative measurement of ThT fluorescence during assembly should reveal a time-dependent increase in intensity that peaks near the time of maximal fibril formation and then declines. This decline, which is commonly observed, results from precipitation of fibrils and sequestration of ThT-reactive sites within supramolecular fibril aggregates. 7.4.2.2 Congo Red Binding Congo red, like ThT, is a dye routinely used to detect β-sheet-rich assemblies.78,79 Upon binding, the absorption maximum of Congo red undergoes a “red shift,” changing from 490 to 540 nm. Because these wavelengths are in the visible range, amyloid binding can be monitored using simple spectrophotometric equipment (see Section 7.4.2.2.1) and optical microscopes (see Section 7.4.2.2.2). In addition, Congo red is birefringent. If macroscopic amyloid assemblies are viewed using polarized light, a yellow–green color is visible in the polarization plane. Cross-polarized light, obtained using two plane polarizers mounted at a 90° angle relative to each other will produce a classic Maltese cross in the presence of amyloid.80

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7.4.2.2.1 Spectrophotometric Monitoring of Congo Red Absorbance 1. Prepare a 20 µM stock solution of Congo red in 20 mM phosphate buffer, pH 7.4, containing 150 µM sodium chloride. 2. Mix 225 µl of the Congo red solution with 25 µl of Aβ. Controls should include Congo red alone and peptide alone. 3. Mix and incubate for 30 min at 22°C. 4. Time-dependent changes in the absorption spectra of samples can be determined by wavelength scans between 400 and 700 nm. Monitoring the red shift of the Congo red intensity maximum is the most reliable indicator of the binding of the dye to amyloid. 5. Quantification of amyloid binding levels can be achieved by measurement of the absorbance of the Congo red:Aβ complex at 541 and 403 nm. The amount of bound Congo red is then calculated according to the following formula, where C is the concentration (µM) of the Congo red:Aβ complex and λA is the total absorbance of the sample at wavelength λ: C=

(

541

) (

A 47,800 –

403

)

A 38,100 .

Note: Amyloid assemblies are quite large and therefore scatter appreciable quantities of light. This scattering reduces the measured light intensity in a wavelength-dependent manner and, depending on the formulae used for quantification, produces artificially high or low estimations of the amount of Congo red bound. At low Aβ concentrations (<5 µg/ml), the error is relatively small. However, at higher concentrations, scattering must be taken into account. Klunk et al. studied these issues extensively and developed protocols for determining absolute amounts of fibrillar Aβ (for a recent review, see Klunk et al.79). These protocols use the absorbance values for the Congo red (ACR) and peptide (AAβ ) controls (step 2 above) to adjust for scattering. The adjustment produces the following formula, useful for Aβ samples with and without significant scattering: Aβ fib =

(

541

)

A 4780 –  403 A ( 4780 r )  +

(

403

ACR (r 4780 ) – (1 3810 ) 

)

where Aβfib is the concentration of fibrillar Aβ in µg/ml, 541A and 403A are the total absorbance values of the sample, 403ACR is the absorbance of the Congo red control, and r = 541AAβ / 403AAβ is the ratio of scattering of the peptide control at 541 and 403 nm. 7.4.2.2.2 Microscopic Monitoring of Congo Red Binding 1. To stain macromolecular aggregates, samples are pelleted by centrifugation at 16,000 × g for 30 min. 2. The resulting pellets then are stained by addition of 100 µl of 20 µM Congo red in phosphate buffer, gentle agitation, and incubation for 5 min at 22°C. 3. Excess Congo red is removed from the labeled aggregates by centrifugation at 16,000 × g for 2 min, removal of the supernate, suspension of the

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pellet in 100 µl of 50% (v/v) ethanol in water, recentrifugation, and repetition of the wash procedure once. 4. The washed pellet is suspended in 50 µl of phosphate buffer and spread evenly on a glass microscope slide. 5. Examination of the sample in a light microscope fitted with crossed polarizers will reveal Maltese cross-like structures if amyloid fibrils are present. Note: It should be noted that fibrillar preparations of Aβ are birefringent regardless of whether they are treated with dyes. For example, at concentrations of 5 to 10 mg/ml in the absence of dyes, striking birefringence can be observed in fibrillar Aβ samples viewed using cross-polarized light.81 However, dye binding provides a more sensitive indicator of this optical property. 7.4.2.3 Turbidity Turbidity, the scattering of light by large particles (scatterers) in solution,82 provides a simple, rapid method to examine Aβ fibril formation.38,49,83,84 Initially, when the peptide exists in a monomeric or oligomeric form, the intensity of scattered light is low and the solution appears clear to the eye. As fibril assembly proceeds, scatterer size increases, producing significant apparent light absorption when samples are monitored spectrophotometrically, and eventually producing a turbid solution with a typically milky appearance. A plot of absorbance versus time will reveal a quasisigmoidal curve. The initial, relatively flat portion of the curve, often called the lag phase, occurs during the period of initial peptide assembly. Scattering intensity varies as the square of molecular weight; therefore, while the peptide aggregates are relatively small, they scatter insufficient light to be detected. As the aggregates grow larger, the intensity of light scattered from them increases to the point that changes in absorbance are observable. At this point the lag phase ends and a rapid increase in absorbance is observed, consistent with the aggregate growth. Eventually the average particle size reaches a maximum, at which point the turbidity remains constant. In samples in which interparticle interactions occur, large fibril masses may precipitate, causing a late decrease in turbidity as these scatterers leave the illuminated area of the sample cuvette. [Readers are cautioned not to equate the term “lag phase,” as associated with turbidity, Congo red, ThT, or similar measures of amyloid fibril formation, with the lag phase associated with nucleation-dependent polymerization reactions. The latter is a measure of the time required for the self-association of monomers to produce fibril nuclei. The lifetimes of nuclei are exceedingly short because they are immediately elongated through monomer addition. Nucleation is not measured directly in the assays mentioned above.] The following protocol is illustrative of the basic strategy for performing turbidity experiments. The precise sample preparation method can be varied to suit the particular experimental needs of the investigator.

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1. Prepare the Aβ sample for study. If gross questions about fibril formation are being studied, one might, for example, prepare lyophilized Aβ peptide in 0.06 M NaOH at a concentration of 4 mg/ml. 2. Add an equal volume of 2× phosphate-buffered saline, pH 7.4, to produce a final peptide concentration of 2 mg/ml. 3. Sonicate the peptide solution for 10 min in a bath sonicator (#1200-R, Branson). 4. Clarify the peptide solution by centrifugation at 16,000 × g for 10 min. 5. Carefully transfer the supernate to a cuvette and place the cuvette in a spectrophotometer. 6. Monitor absorbance at 400 nm. Note: Rates of fibril assembly can vary dramatically, depending on the peptide studied and the conditions of the assembly reaction. Aggregate-free preparations can take weeks or longer [one Aβ solution has remained monomeric for years (J. Lee, personal communication)] to assemble. Agitation, a pH near the pI of Aβ (5.5), increased salt concentration, increased temperature, and other manipulations can accelerate the assembly kinetics.

7.4.3 SECONDARY STRUCTURE DETERMINATION Amyloid formation involves reorganization of secondary structure elements within the amyloid protein monomer. This is a prerequisite for assembly of the classic extended sheet structures comprising amyloid fibrils.10,11 The reorganization process occurs in two ways, via organization of initially unstructured peptide (e.g., α-synuclein or synthetic Aβ) or destabilization of initially structured protein (e.g., transthyretin or prion). During Aβ fibril formation, the unstructured monomer forms a partiallystructured intermediate containing a significant amount of helix.18 This intermediate then undergoes a conformational transition to the β-sheet-rich form present in fibrils. Monitoring peptide secondary structures provides the means to study basic features of amyloid assembly and how changes in primary structure, solvent conditions and the presence of other proteins or chemicals affect amyloidogenesis. CD spectroscopy (Section 7.4.3.1) and Fourier transform infrared spectroscopy (FTIR; Section 7.4.3.2) are two common methods for monitoring protein secondary structure. 7.4.3.1 CD Spectroscopy Circular dichroism (CD) is the difference in the absorption of left and right circularly polarized light. Optically active samples exhibit CD, and thus the acquisition and analysis of CD data allow inferences to be made about the structures within samples. In proteins, secondary structure elements, including α-helices, β-strands, β-turns, and disordered regions display characteristic wavelength-dependent CD. CD thus is a useful tool for determining protein secondary structure content. CD also is used frequently to determine the stability of the folded states of proteins by monitoring the levels of α-helix or β-sheet during variations in temperature

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or in the concentrations of chemical denaturants (usually urea or guanidinium hydrochloride (Gu·HCl)). The simplest experiment monitors the two-state conformational folded↔unfolded transition. For example, temperature and Gu · HCl denaturation–renaturation experiments have been used to study the effects of TFE on Aβ40 and Aβ42 helix stability and fibrillogenesis kinetics.85 1. Prepare LMW Aβ at a concentration of ~30 µM in 10 mM sodium phosphate, pH 7.4. 2. Incubate the sample under the desired experimental conditions (see note below). 3. Transfer the sample into a quartz CD cuvette. A 0.1-cm path-length quartz cell (Hellma, Forest Hills, NY) is a useful compromise between the necessities (due to generally low peptide concentrations) of having the longest path length possible and the lowest solvent absorption possible. If the peptide sample is not maintained within the CD cuvette during the experiment, transfers to and from the cuvette should be made as gently as possible (especially under nonagitated incubation conditions). 4. CD measurements can be performed on any standard spectropolarimeter, according to the manufacturer’s instructions. Jasco, Inc. (Easton, MD) and Aviv Associates (Lakewood, NJ) both supply suitable instruments. Measurements should be made at the same temperature used to incubate the sample. Greater accuracy in spectral interpretation is obtained if a lower wavelength limit of 185 to 190 nm can be achieved, although for many applications this is not essential, and for some, not obtainable. Multiple spectra should be acquired in order to increase the signal-tonoise ratio and obtain smooth spectra. 5. A CD spectrum from buffer alone should always be acquired and subtracted from the sample spectra. 6. Raw spectra typically are acquired as tables of absolute ellipticity vs. wavelength. To normalize for peptide concentration, it is preferable to determine the mean residue ellipticity, [Θ]λ (deg cm2 dmol–1), according to the formula [Θ]λ = M Θλ /10dcn, where M is peptide molecular weight (g/mol), Θλ is ellipticity (degrees), d is path length (cm), c is peptide concentration (g/ml), and n is the number of amino acids in the peptide. The resulting spectra then can be interpreted both qualitatively and quantitatively through comparison with spectra from standard proteins or peptides. A variety of spectral deconvolution packages are available (e.g., CDANAL86 and CDPro87) that will produce quantitative estimates of the levels of specific secondary structure elements, including random coil, α-helix, β-sheet, and β-turn. Readers are cautioned, however, that none of these methods is entirely accurate and therefore the numbers they produce must be considered approximations. Note: Sample preparation method, buffer composition, incubation conditions, and CD acquisition parameters all may be varied to suit experimental needs. However, a number of factors must be considered. Increased sensitivity is obtained by using more concentrated peptide solutions. Buffers should be UV-transparent so that

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a wavelength range of sufficient magnitude can be scanned. Temperature must be controlled carefully so that variations do not occur during sample manipulation outside of and inside the CD instrument. 7.4.3.2 Fourier Transform Infrared Spectroscopy (FTIR) Infrared spectroscopy is a well-established optical method for determining protein secondary structures.88–91 FTIR provides the means to determine protein conformation within samples in multiple milieus, including in solution, in thin films, and as solids. The technique is based on the fact that absorption spectra of peptides and proteins acquired in the infrared wavenumber range (1600 to 1700 cm-1) produce peaks characteristic of specific secondary structure elements.89–95 These elements include α-helix, 310-helix, extended chain/β-sheet, loop, β-turn, aggregated strands, antiparallel β-sheet, and disordered (random coil). Characteristic differences have been reported between β-sheet structures in monomeric and aggregated species. The FTIR method has been used quite successfully for secondary structure determinations of Aβ samples.31.89,96

7.4.4 TOPOGRAPHICAL ANALYSIS Protein folding in general, and amyloid protein assembly in particular involve the formation of tertiary and quaternary structures in which amino acids can be considered exposed or buried with respect to solvent water. The packing of buried residues is involved in the nucleation of intramolecular protein folding and the organization and stabilization of folded structures both in protein monomers and large assemblies. Monitoring the topography of an amyloid protein provides important insights into the mechanisms of its folding and assembly. 7.4.4.1 8-Anilino-1-Naphthalenesulfonic Acid (ANS) Binding ANS, a fluorescent dye, binds to solvent-exposed hydrophobic surfaces, such as those found in partially folded intermediate (molten globule) states.97 When bound, an 8- to 10-fold increase in ANS fluorescence intensity occurs. This large increase, along with the rather weak affinity ANS has for native or completely unfolded states, makes it a useful probe of protein folding. This is true for amyloid assembly as well, which involves the formation of partially folded (e.g., α-synuclein and Aβ) and partially unfolded (e.g., transthyretin and prion) intermediates. 1. Prepare stock 100 µM ANS in 10 mM phosphate buffer, pH 7.5. 2. Mix 50 µl of an Aβ sample with 50 µl of the ANS stock solution. 3. Measure the fluorescence intensity immediately after mixing. ANS should be excited at a wavelength of 370 nm. Its emission can be measured within the wavelength range of 400 to 500 nm. Fluorometers are available commercially from many sources. We use an F4500 spectrofluorometer (Hitachi Instruments, Rye, NH). 4. Binding of ANS to hydrophobic patches causes a blue wavelength shift (toward lower wavelengths) for the fluorescence intensity maximum along with an absolute increase in the fluorescence intensity.

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Note: The Aβ sample can be studied in phosphate buffer or alternate buffers can be used. In the latter case, the investigator should ensure that the fluorescence properties of the dye are unaffected. 7.4.4.2 Intrinsic Fluorescence The fluorescence characteristics of the aromatic amino acids tryptophan (Trp), tyrosine (Tyr), and phenylalanine (Phe) provide the means to monitor the conformational dynamics of protein folding and assembly.98 Trp is a particularly attractive amino acid for use in intrinsic fluorescence experiments because it fluoresces strongly and the emission spectrum of the Trp indole ring is highly sensitive to solvent polarity and to the presence of fluorescence quenchers. Changes in the immediate vicinity of Trp that occur during protein folding and assembly thus are indicated clearly by alterations in Trp fluorescence. Tyr can also be used to probe local environmental changes, although it fluoresces less strongly than Trp. Tyr fluorescence (λmax), in contrast to that of Trp, is almost insensitive to solvent polarity. However, Tyr fluorescence can be quenched by exposure of the phenol ring to hydrated peptide carbonyl groups. Quenching also may be produced through hydrogen bond formation with peptide carbonyls or with carboxylate side chains of aspartic or glutamic acid. Intrinsic tyrosine fluorescence thus reveals features of the local environment of the phenol group. Aβ40 and Aβ42 contain a single Tyr at position 10; thus intrinsic fluorescence experiments can be done on wild-type peptides without modification to their primary structure. 1. Transfer 100 µl of Aβ solution into a rectangular 10-mm quartz microcuvette. The fluorescence intensity of Tyr10 in 50 to 100 µM wild-type Aβ is substantial (10,000 arbitrary units), thus Aβ assembly reactions carried out in the micromolar concentration regime are amenable to study by this method. We routinely use 50 µM sodium phosphate, pH 7.5, containing 0.01% (w/v) sodium azide as the buffer in this method. 2. Place the cuvette into a fluorometer and measure the fluorescence using an excitation wavelength of 280 nm. Spectra can be acquired by scanning from 290 to 500 nm. For assay purposes, fluorescence at a wavelength of 303 nm can be monitored. Slit widths used for excitation and emission are 5 and 10 nm, respectively, with a scan rate of 240 nm/min. 3. The fluorescence emission spectrum of phosphate buffer (background intensity) is subtracted from the emission spectrum of the Aβ samples. 7.4.4.3 Electron Paramagnetic Resonance (EPR) EPR is a technique for monitoring changes in the spin state of electrons. It can be applied to free radicals and other molecular species that possess unpaired electrons. Alterations in spin state reveal molecular features of the chemical environment, structure, and motional dynamics of the electrons under study. Interactions between two spin systems can also provide distance information. EPR is a powerful approach for probing the dynamics of protein folding and assembly.99–105 Typically, a spin label (a stable free radical such as a nitroxide) is

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covalently attached to a specific site in a protein. Cysteine residues, either present natively in the protein or placed at precise locations through genetic engineering, are convenient sites for attachment. Analysis of EPR spectra during the assembly process allows correlation of local environmental features around the spin label with the global structural organization of the protein assembly. Factors controlling the EPR spectrum include the solvent-accessibility of the probe and the motional freedom of the spin-labeled side chain. It has been shown that backbone secondary structure and aspects of the tertiary structure can be determined from periodic variations in the accessibility of nitroxide side chains in a sequential set of spin-labeled proteins.100 It also has been established that the motional dynamics of the side chain reflected in the EPR spectral line shape correlates with general features of the protein fold.100 An excellent example of the use of EPR for understanding the structural organization of Aβ in amyloid fibrils comes from the work of Langen et al. who used site-directed spin labeling (SDSL).106 Nineteen different spin-labeled Aβ peptides were synthesized and each was allowed to form fibrils. Multiple techniques were used to demonstrate that the spin label did not alter the ability of each peptide to fold and assemble into fibrils. Information from spin–spin interactions between equivalent residues in different peptides revealed that the fibrils formed by both Aβ40 and Aβ42 have arrangements of their peptide backbones that are parallel and inregister. Motional analysis showed that the central region of Aβ, residues 14-38, is relatively stable, whereas significant mobility exists at both the N and C termini. Interestingly, in contrast to some amyloids, Aβ40 and Aβ42 formed mixed fibrils upon coincubation. The SDSL EPR technique is a general method. For example, it has been used to study transthyretin amyloid formation.107 7.4.4.4 Hydrogen–Deuterium Exchange Hydrogen exchange is a phenomenon in which covalently bound, but labile, hydrogen atoms of proteins exchange (switch places) at a finite rate with solvent hydrogen atoms.108 Hydrogen exchange is a powerful probe of protein structure and dynamics. The technique typically involves exchange of labile protein hydrogens with deuterons from solvent 2H2O (D2O) and thus is called hydrogen–deuterium exchange. Labile hydrogens in proteins are those bonded to nitrogen, sulfur or oxygen. The exchange of backbone amide hydrogens is studied frequently because their exchange rates are amenable to monitoring and they play key roles in hydrogen bonding interactions in many structural elements of proteins (e.g., α-helices and β-strands). For a backbone amide hydrogen to exchange with a solvent deuteron, the amide hydrogen must be solvent-exposed and not involved in hydrogen bonding. These requirements make hydrogen exchange sensitive to structural rearrangements that occur during protein aggregation. As oligomeric and higher-order assemblies form, the amide hydrogens buried in the core of the assembly or hydrogen bonded in α-helices and β-sheets do not exchange readily with solvent deuterons. This phenomenon is known as protection. Kheterpal and coworkers have shown that 40% of the backbone amide hydrogens of protofibrils formed by Aβ40 are protected from exchange.60 In contrast, 60% of the backbone amide hydrogens are protected in mature amyloid fibrils.109

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A typical hydrogen exchange experiment involves three main steps: (1) formation and isolation of the assembly of interest; (2) exposure of the assembly to an excess of D2O; and (3) detection of exchange. Detection techniques include NMR and mass spectrometry (MS). NMR can pinpoint at the residue level the site where the exchange took place. However, it is limited by the need for high, nonphysiological protein concentrations and by a molecular mass limit of ~35 kDa. MS can measure the increase in mass that results from the exchange of hydrogens (1 amu/hydrogen exchanged). Advantages of MS include lower (by orders of magnitude) required concentrations of protein and applicability to very high mass protein assemblies and complexes. Furthermore, following endoproteolysis with the acid protease pepsin, MS experiments can be used to identify peptides in which hydrogen exchange occurred and then to systematically fragment them to determine precisely which amides underwent exchange. In-depth discussions of theoretical and practical aspects of the use of NMR and MS to monitor hydrogen exchange may be found in recent reviews by Engen and Smith110 and Dempsey.111

7.4.5 NMR SPECTROSCOPY Solution-state NMR is an extremely powerful technique for determining high-resolution structures of stable protein conformers and monitoring protein dynamics. NMR generally is performed at relatively high protein concentrations (1 mM) and requires that the protein of interest be soluble and unaggregated. Because of the propensity of Aβ to aggregate, these requirements have complicated NMR studies of this peptide under quasiphysiologic conditions. NMR studies of full-length Aβ have been conducted in the presence of high concentrations of organic cosolvents, including HFIP112 and TFE,113 or detergents (e.g., sodium dodecyl sulfate).114 Under these conditions, aggregation is blocked but formation of α-helices is facilitated significantly. Studies done using these solvent conditions thus have yielded insights into α-helix formation and stabilization. An alternate approach to controlling Aβ selfassociation has been to conduct NMR experiments at low temperature (e.g., 5°C) or to use Aβ in which Met35 has been oxidized to its sulfoxide form.115 This modification has been shown to delay the oligomerization of Aβ.116,117 Results from these studies have revealed a predominantly random, extended-chain conformation for Aβ40, Aβ42, Met35(O)Aβ40 and Met35(O)Aβ42, and turn- or bend-like structures located at Asp7–Glu11 and Asp23–Ser26.117 One of the most exciting advances in NMR-based studies of Aβ assembly has come from studies of Aβ40 fibrils in the solid state.118 These studies were made possible by improved technology (e.g., probes that allow rotation at speeds up to ~30 kHz) and experiments that permit accurate measurements of intramolecular and intermolecular distances, backbone torsion angles, and chemical shifts that correlate with secondary structures.119 A structural model of Aβ40 protofilaments, derived by energy minimization with constraints based on solid-state NMR data, has been shown to be consistent with the overall dimensions and linear density derived from EM data.118

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7.4.6 MORPHOLOGICAL ANALYSIS In some ways, the fine-structure analyses discussed to this point may be likened to the proverbial blind men describing an elephant.120 Each technique provides important information about a particular peptide subregion or assembly process, but does not provide a global view of the peptide assembly itself. In the following two sections, we discuss in general terms the use of electron microscopy (EM) and atomic force microscopy (AFM) for determining the morphologies of Aβ assemblies. 7.4.6.1 Electron Microscopy Electron microscopy has been used extensively to examine the morphology of Aβ assemblies. In one approach, the protein sample is spotted onto a small support (grid), allowed to adhere and then washed briefly with water. The assemblies thus immobilized are chemically cross-linked with glutaraldehyde to stabilize them and then stained with an electron-dense dye such as uranyl acetate or phosphotungstic acid. The dye provides contrast so that the assemblies can be visualized after placement inside the electron microscope. The images obtained show dark structures against a white background; thus this technique is referred to as negative staining. In addition to direct examination of negatively stained preparations, metal casts of the assemblies can be constructed using a technique called rotary shadowing.121 This produces very high contrast images. Care must be exercised in the conduct and interpretation of EM studies because artifactual results are easy to produce. These artifacts arise for numerous reasons. Particulates in the EM stains can be interpreted as proteinaceous. Damage to the grid surface may be interpreted as protein sheets. Specific subsets of assemblies present in a protein sample may adhere to the grid, whereas others may not. This leads to erroneous conclusions about the distributions of morphologies present. Even if all assemblies adhere, because only a small area of a grid is illuminated at any one time, it is critical to sample a sufficient number of locations to ensure that a representative sampling of the bound assemblies is obtained. A substantial literature exists concerning EM studies of Aβ. The assemblies examined include LMW Aβ,17,116 small oligomers,122–124 paranuclei,17,116 protofibrils,42,60,61,66,122,124 spheroids,26,27,125 and fibrils.24,38,66,121,125,126 7.4.6.2 Atomic Force Microscopy AFM is a method for morphological analysis that complements EM. AFM provides three-dimensional images of assemblies adherent to an atomically flat support, typically mica. AFM is a physical method in which a small probe is rastered across a mica surface. The probe either remains in continuous contact with the surface or taps the surface at high frequency. As the probe encounters discontinuities in the surface due to the presence of adherent material, its vertical position changes and these changes are recorded by the instrument. Thus, after complete scanning of the AFM support, a relief map can be constructed in which extremely fine (nanometer) detail can be revealed.

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The resolution of this method is limited by the size and shape of the probe tip. Therefore, although the heights of objects can be measured accurately, widths can be overestimated.25 A powerful feature of AFM that cannot be achieved by EM is the continuous monitoring of protein assembly in the “solution phase.” Experiments can be done in which fibril growth occurs on an AFM support immersed in solvent.127 This allows real-time monitoring of assembly growth and kinetics. Although obvious, it is important to appreciate that any AFM method can only reveal features of material that will adsorb to the mica surface. Thus, as with EM, conclusions about population morphologies must be made cautiously. AFM has been used extensively to study the morphology of Aβ oligomers,20,33,128 protofibrils,12,128 and fibrils.128–131

7.5 DISCUSSION Since the time of Virchow, when amyloid was thought to be a complex carbohydrate (and thus the misnomer “amyloid” was introduced), the study of amyloid protein structure, assembly and biological activity has been problematic. To a large degree, this is because of the strong propensity of amyloid proteins to self-associate, which makes the use of classical methods in structural biology either difficult or impossible. Aβ, in particular, has been a vexing subject of study because the native monomeric state of this peptide remains largely undefined and the peptide exists in equilibrium with dimers and larger oligomers. Therefore, unlike proteins with well-defined native folds, Aβ folding and assembly proceeds from a poorly defined starting conformer to an incompletely defined amyloid fibril. Nevertheless, significant progress has been made in elucidating pathways of Aβ assembly. In this chapter, we have sought to provide both the theoretical and practical tools necessary to study this enigmatic peptide. We also have pointed out pitfalls that must be fully understood and considered in experimental planning if valid and useful conclusions are to be derived from studies of Aβ. In this regard, the following maxim of Sir Isaac Newton (1642–1727) may have special applicability for studying amyloid protein assemblies: To explain all nature is too difficult a task for any one man or even for any one age. ’Tis much better to do a little with certainty, and leave the rest for others that come after you, than to explain all things.

ACKNOWLEDGMENTS We thank Dr. Gal Bitan and Ms. Sabrina Vollers for critical comments. We gratefully acknowledge the support of the National Institutes of Health (NS38328, AG18921, and NS44147) and the Foundation for Neurologic Diseases.

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67. Tomski, S.J. and Murphy, R.M., Kinetics of aggregation of synthetic β-amyloid peptide, Arch. Biochem. Biophys. 294, 630–638, 1992. 68. Shen, C.L. et al., Light scattering analysis of fibril growth from the amino-terminal fragment β(1-28) of β-amyloid peptide, Biophys. J. 65, 2383–2395, 1993. 69. Murphy, R.M. and Pallitto, M.R., Probing the kinetics of β-amyloid self-association, J. Struct. Biol. 130, 109–122, 2000. 70. Lomakin, A. et al., Kinetic theory of fibrillogenesis of amyloid β-protein, Proc. Natl. Acad. Sci. USA 94, 7942–7947, 1997. 71. Yong, W. et al., Structure determination of micelle-like intermediates in amyloid β-protein fibril assembly by using small angle neutron scattering, Proc. Natl. Acad. Sci. USA 99, 150–154, 2002. 72. Kusumoto, Y. et al., Temperature dependence of amyloid β-protein fibrillization, Proc. Natl. Acad. Sci. USA 95, 12277–12282, 1998. 73. Lomakin, A., Benedek, G.B., and Teplow, D.B., Monitoring protein assembly using quasielastic light scattering spectroscopy, Methods Enzymol. 309, 429–459, 1999. 74. Rogers, D.R., Screening for amyloid with the thioflavin-T fluorescent method, Am. J. Clin. Pathol. 44, 59–61, 1965. 75. LeVine, H., III, Quantification of β-sheet amyloid fibril structures with thioflavin T, Methods Enzymol. 309, 274–284, 1999. 76. LeVine, H., III, Thioflavin T interaction with synthetic Alzheimer’s disease β-amyloid peptides: detection of amyloid aggregation in solution, Prot. Sci. 2, 404–410, 1993. 77. Naiki, H. and Nakakuki, K., First-order kinetic model of Alzheimer’s β-amyloid fibril extension in vitro, Lab. Invest. 74, 374–383, 1996. 78. Klunk, W.E., Pettegrew, J.W., and Abraham, D.J., Quantitative evaluation of Congo red binding to amyloid-like proteins with a β-pleated sheet conformation, J. Histochem. Cytochem. 37, 1273–1281, 1989. 79. Klunk, W.E., Jacob, R.F., and Mason, R.P., Quantifying amyloid by Congo red spectral shift assay, Methods Enzymol. 309, 285–305, 1999. 80. Westermark, G.T., Johnson, K.H., and Westermark, P., Staining methods for identification of amyloid in tissue, Methods Enzymol. 309, 3–25, 1999. 81. Malinchik, S.B. et al., Structural analysis of Alzheimer’s β(1-40) amyloid: protofilament assembly of tubular fibrils, Biophys. J. 74, 537–545, 1998. 82. Andreu, J.M. and Timasheff, S.N., The measurement of cooperative protein selfassembly by turbidity and other techniques, Methods Enzymol. 130, 47–59, 1986. 83. Jarrett, J.T., Berger, E.P., and Lansbury, P.T., Jr., The C-terminus of the β protein is critical in amyloidogenesis, Ann. N. Y. Acad. Sci. 695, 144–148, 1993. 84. Evans, K.C. et al., Apolipoprotein E is a kinetic but not a thermodynamic inhibitor of amyloid formation: implications for the pathogenesis and treatment of Alzheimer’s disease, Proc. Natl. Acad. Sci. USA 92, 763–767, 1995. 85. Fezoui, Y. and Teplow, D.B., Kinetic studies of amyloid β-protein fibril assembly: differential effects of α-helix stabilization, J. Biol. Chem. 277, 36948–36954, 2002. 86. Perczel, A., Park, K., and Fasman, G.D., Analysis of the circular dichroism spectrum of proteins using the convex constraint algorithm: a practical guide, Anal. Biochem. 203, 83–93, 1992. 87. Sreerama, N., Venyaminov, S.Y., and Woody, R.W., Estimation of protein secondary structure from circular dichroism spectra: inclusion of denatured proteins with native proteins in the analysis, Anal. Biochem. 287, 243–251, 2000. 88. Jackson, M. and Mantsch, H.H., The use and misuse of FTIR spectroscopy in the determination of protein structure, Crit. Rev. Biochem. Mol. Biol. 30, 95–120, 1995.

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112. Crescenzi, O. et al., Solution structure of the Alzheimer amyloid β-peptide (1-42) in an apolar microenvironment: similarity with a virus fusion domain, Eur. J. Biochem. 269, 5642–5648, 2002. 113. Sticht, H. et al., Structure of amyloid A4-(1-40)-peptide of Alzheimer’s disease, Eur. J. Biochem. 233, 293–298, 1995. 114. Shao, H.Y. et al., Solution structures of micelle-bound amyloid β-(1-40) and β-(1-42) peptides of Alzheimer’s disease, J. Mol. Biol. 285, 755–773, 1999. 115. Riek, R. et al., NMR studies in aqueous solution fail to identify significant conformational differences between the monomeric forms of two Alzheimer peptides with widely different plaque-competence, Aβ(1-40)ox and Aβ(1-42)ox, Eur. J. Biochem. 268, 5930–5936, 2001. 116. Bitan, G. et al., A molecular switch in amyloid assembly: Met35 and amyloid β-protein oligomerization, J. Am. Chem. Soc. 125, 15359–15365, 2003. 117. Hou, L. et al., Solution NMR studies of the Aβ(1-40) and Aβ(1-42) peptides establish that the Met35 oxidation state affects the mechanism of amyloid formation, J. Am. Chem. Soc. 126, 1992 –2005, 2004. 118. Tycko, R., Progress towards a molecular-level structural understanding of amyloid fibrils, Curr. Opin. Struct. Biol. 14, 1–8, 2004. 119. Tycko, R., Insights into the amyloid folding problem from solid-state NMR, Biochemistry 42, 3151–3159, 2003. 120. Blind men and the elephant, http://swaminarayansatsang.com/literature/literaturestory.asp?StoryID=26. 121. Goldsbury, C.S. et al., Studies on the in vitro assembly of Aβ 1-40: implications for the search for Aβ fibril formation inhibitors, J. Struct. Biol. 130, 217–231, 2000. 122. Nybo, M., Svehag, S.E., and Nielsen, E.H., An ultrastructural study of amyloid intermediates in Aβ1-42 fibrillogenesis, Scand. J. Immunol. 49, 219–223, 1999. 123. Roher, A.E. et al., Morphology and toxicity of Aβ(1-42) dimer derived from neuritic and vascular amyloid deposits of Alzheimer’s disease, J. Biol. Chem. 271, 20631–20635, 1996. 124. Modler, A.J. et al., Assembly of amyloid protofibrils via critical oligomers: a novel pathway of amyloid formation, J. Mol. Biol. 325, 135–148, 2003. 125. Antzutkin, O.N., Amyloidosis of Alzheimer’s Aβ peptides: solid-state nuclear magnetic resonance, electron paramagnetic resonance, transmission electron microscopy, scanning transmission electron microscopy and atomic force microscopy studies, Magn. Reson. Chem. 42, 231–246, 2004. 126. Tjernberg, L.O. et al., A molecular model of Alzheimer amyloid β-peptide fibril formation, J. Biol. Chem. 274, 12619–12625, 1999. 127. Blackley, H.K.L. et al., In situ atomic force microscopy study of β-amyloid fibrillization, J. Mol. Biol. 298, 833–840, 2000. 128. Legleiter, J. et al., Effect of different anti-Aβ antibodies on Aβ fibrillogenesis as assessed by atomic force microscopy, J. Mol. Biol. 335, 997–1006, 2004. 129. Harper, J.D., Lieber, C.M., and Lansbury, P.T., Atomic force microscopic imaging of seeded fibril formation and fibril branching by the Alzheimer’s disease amyloid-β protein, Chem. Biol. 4, 951–959, 1997. 130. Stine, W.B., Jr. et al., The nanometer-scale structure of amyloid-β visualized by atomic force microscopy, J. Protein Chem. 15, 193–203, 1996. 131. Kowalewski, T. and Holtzman, D.M., In situ atomic force microscopy study of Alzheimer’s β-amyloid peptide on different substrates: new insights into mechanism of β-sheet formation, Proc. Natl. Acad. Sci. USA 96, 3688–3693, 1999.

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8

Intracellular Accumulation of Amyloid β and Mitochondrial Dysfunction in Down’s Syndrome Jorge Busciglio, Alejandra Pelsman, Pablo Helguera, and Atul Deshpande

CONTENTS Abstract 8.1 Introduction 8.2 Experimental Procedures 8.2.1 Cell Culture 8.2.2 Antibodies 8.2.3 Immunocytochemical Procedures 8.2.3.1 Extraction 8.2.3.2 Fixation 8.2.3.3 Second Labeling Protocol 8.2.4 Western Blot. 8.2.5 Inhibition of Energy Metabolism 8.2.6 Assessment of Mitochondrial Function 8.2.6.1 MTS Assay 8.2.6.2 JC1 Protocol 8.2.7 Cell Viability Assays 8.2.7.1 Trypan Blue 8.2.7.2 Propidium Iodide 8.2.8 Neuroprotection Assays 8.3 Results 8.3.1 Detection of Intracellular Aβ in DS Astrocytes 8.3.2 DS-Like Alterations in APP Processing Induced in Normal Astrocytes by Energy Depletion 8.3.3 Mitochondrial Dysfunction in DS Astrocytes 8.3.4 APPs Rescues DS Cortical Neurons from Apoptosis 8.4 Discussion Acknowledgments References 0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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ABSTRACT Cortical astrocyte and neuronal cultures derived from fetal Down’s syndrome (DS) brain were utilized to demonstrate intracellular accumulation of insoluble Aβ42 and inhibition of mitochondrial energy metabolism consistent with impaired mitochondrial function in DS. Similarly, energy depletion in normal astrocytes resulted in intracellular Aβ accumulation, suggesting an important role for mitochondrial dysfunction in the generation and accumulation of intracellular Aβ. Energy deficits in DS cells also alter amyloid precursor protein (APP) processing, resulting in decreased APP secretion in vitro and in vivo. The survival of DS cortical neurons was markedly increased by addition of recombinant APP or conditioned medium of normal astrocyte cultures, but not by conditioned medium depleted of secreted APP (APPs), suggesting that APPs may be a survival factor for human neurons. These results indicate that mitochondrial dysfunction in DS brain cells leads to intracellular deposition of Aβ42, reduced levels of APPs, and a chronic state of increased neuronal vulnerability.

8.1 INTRODUCTION Down’s syndrome (DS) or trisomy 21 is the most common genetic cause of mental retardation. The neuropathology of DS is complex and includes decreased brain weight and neuronal number, abnormal neuronal differentiation, and structural changes in dendritic spines.1 A distinct feature of DS is the onset of Alzheimer’s disease (AD) by middle age.1,2 The development of AD in DS may be related to overexpression of the amyloid precursor protein (APP) due to increased gene dosage, leading to increased Aβ generation.3,4 Deficits in mitochondrial function cause selective neuronal degeneration and may be involved in a number of neurodegenerative disorders.5,6 DS cortical neurons in culture exhibit intracellular accumulation of reactive oxygen species (ROS) and increased lipid peroxidation leading to neuronal apoptosis.7 Interestingly, perturbations of mitochondrial homeostasis constitute an early and critical feature of apoptotic processes that precede free radical formation and neuronal death.8 Energy depletion and oxidative stress can also induce amyloidogenic changes in APP processing,9 suggesting a potential link between mitochondrial dysfunction, oxidative stress, and Aβ production. In this regard, dysfunction of mitochondrial electron transport proteins and a close relationship between mitochondrial abnormalities and oxidative damage have been described in AD brains.10–12 To characterize the molecular events involved in DS neuropathology and the development of AD in DS, we analyzed Aβ generation and mitochondrial activity in normal and DS cortical astrocytes in culture and in the DS brain. The results suggest that mitochondrial dysfunction may play a significant role in the development of AD neuropathology in DS patients by promoting aberrant APP processing and intracellular accumulation of Aβ. Below is a detailed description of the techniques utilized for immunodetection of intracelllular Aβ and for the assessment of mitochondrial function and cell viability in normal and DS cortical astrocyte and neuronal cultures.

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8.2 EXPERIMENTAL PROCEDURES 8.2.1 CELL CULTURE Primary human cortical cultures were established from the cerebral cortices of normal and DS-aborted fetuses from embryonic weeks 17 through 21. Twelve DS brain specimens and 12 age-matched controls were used to generate neuronal and astrocyte cultures. The protocol for tissue procurement complied with federal and institutional guidelines for fetal research. Cortical astrocyte cultures were prepared as previously described.13 Cells were plated on culture dishes or glass coverslips at a density of 100,000 cells/cm2 and maintained in 10% iron-supplemented calf serum (HyClone, Logan, UT) and D-MEM (Life Technologies, Invitrogen, Carlsbad, CA). For all experiments, astrocyte cultures were fixed or harvested between 30 and 35 days in vitro. More than 95% of the cells present in the cultures stained positive for glial fibrillary acidic protein (GFAP). Normal and DS cortical neuronal cultures were prepared as previously described.7,14

8.2.2 ANTIBODIES C8 is a polyclonal antibody directed against residues 676–695 of APP which recognizes holo-APP and C-terminal fragments. Alz90 (Roche, Basel, Switzerland) and 8E15 (supplied by Dr. Peter Seubert, Elan Pharmaceuticals, Dublin, Ireland) are monoclonal antibodies that recognize residues 511–608 and 444–592, respectively, and recognize APPs. Monoclonal antibodies α-Aβ42 and α-Aβ40 specifically recognize the free C terminus of Aβx-42 and Aβx-40, respectively.16,17 Other antibodies used included anti-syntaxin 6, anti-early endosomal antigen 1 (Signal Transduction Laboratories, BD Biosciences, Canada), anti-LAMP1 (Signal Transduction Laboratories), mouse anti-α-tubulin and anti-α-tubulin isotype III (Sigma Chemical, St. Louis, MO, USA), and mouse anti-Cu/Zn superoxide dismutase (Sigma).

8.2.3 IMMUNOCYTOCHEMICAL PROCEDURES Double immunofluorescence was performed on fixed cultures using two consecutive protocols (Figure 8.1). In the first step, intracellular Aβ in cultured astrocytes was immunolabeled using a tyramide signal amplification kit (TSA, NEN Life Science Products, now PerkinElmer, Boston, MA). This technique is designed to enhance signal labeling approximately 1000-fold over regular protocols using biotinylated secondary antibodies. The TSA indirect assay uses the capability of horseradish peroxidase (HPR) to generate reactive tyramide radicals with very short half-lives, resulting in biotinyl–tyramide depositions very close to the sites of enzymatic activities. Biotin is detected by streptavidine–fluorophore conjugates, in this case streptavidine–Oregon green. In the second step, organelle-specific markers were immunolabelled by standard procedures. For some experiments, the cultures were extracted prior to fixation.7 The purpose of this step is to extract soluble proteins while preserving fibrillar and/or insoluble cellular elements in the preparation. The procedure is as follows.

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FIGURE 8.1 Flowchart of general procedure utilized for double immunofluorescence of intracellular Aβ and organelle markers in cultured astrocytes.

8.2.3.1 Extraction 1. Wash cultures with prewarmed stabilizing buffer (0.13 M HEPES, pH 6.9, 2 mM MgCl2, and 10 mM EGTA). 2. Extract cultures in the same buffer plus 0.2% Triton X-100 for 5 min at 37˚C. 3. The cultures are fixed and immunocytochemistry is performed on the TX-100-resistant material remaining on the coverslip. 8.2.3.2 Fixation 1. Fix cultured cells in 4% paraformaldehyde and 0.12 M sucrose in PBS for 20 min at 37˚C. 2. Wash in PBS 3X 5 min each. 3. If a permeabilization step has not been performed prior to fixation, permeabilize with 0.2% Triton X-100 in PBS for 5 min at room temperature. 3. The TSA procedure is performed according to manufacturer’s instructions (refer to NEN Life Sciences TSA-Indirect technical brochure).

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8.2.3.3 Second Labeling Protocol 1. 2. 3. 4.

Block for 1 hr in 5% BSA/PBS. Incubate overnight at 4˚C with corresponding primary antibody. Wash with PBS 0.1% Triton X-100 3X 5 min each. Incubate in Cy3-conjugated mouse or rabbit secondary antibody in 1% BSA/PBS for 30 min at room temperature. 5. Wash with PBS 0.1% Triton X-100 3X 5 min each. 6. Mount coverslips with antifade mounting medium. Fluorescence was visualized with an Olympus IX-70 inverted microscope or a Zeiss LSM 410 confocal-scanning microscope. Specificity was confirmed by preabsorption of the primary antibody with synthetic Aβ1-42 peptide which abolished immunoreactivity.

8.2.4 WESTERN BLOT Cultures were washed with PBS and harvested in RIPA buffer at 4oC. The lysates were centrifuged at 100,000× g for 60 min. Supernatants were mixed 1:1 with SDSreducing sample buffer and boiled for 5 min, followed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). The proteins were electrotransferred to polyvinylidene difluoride membranes, blocked, incubated overnight at 4oC with primary antibody and developed by enhanced chemiluminescence. To control for protein loading, tubulin levels were analyzed in all samples with anti-α-tubulin. Quantitative analysis was performed using a FastScan densitometer (Molecular Dynamics, Sunnyvale, CA) as described.18 Briefly, a standard curve of pixel values was constructed by immunoblotting a serial dilution of purified tubulin. Volume analysis on the appropriate bands was performed using NIH Image software. All densitometric values used for analysis were within the linear range of pixel values.

8.2.5 INHIBITION

OF

ENERGY METABOLISM

To inhibit mitochondrial energy metabolism, cultures were incubated with carbonyl cyanide phenylhydrazone (CCCP). CCCP is one of the most potent uncouplers of both oxidative phosphorylation and photophosphorylation. It produces strong blocking effects on adenosine triphosphate (ATP) generation in mitochondria and chloroplasts.19 Normal astrocytes were incubated with 80 µM of CCCP for 4 hr before harvesting for biochemical analysis. For immunofluorescence microscopy of intracellular Aβ, astrocytes were incubated with 20 µM CCCP for 3 days and with 80 µM CCCP for 4 hr before fixation. This longer incubation protocol does not reduce astrocyte survival as determined by propidium iodide exclusion assay. The longer incubation increases intracellular Aβ levels, enhancing detection by immunocytochemistry or Western blot.

8.2.6 ASSESSMENT

OF

MITOCHONDRIAL FUNCTION

Mitochondrial redox activity was analyzed in normal and DS astrocyte cultures using 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-

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tetrazole (MTS) salt and MTT reduction assays following the vendor’s protocol (Promega, Madison, WI). Both are colorimetric assays that measure the conversion of a tetrazolium component (MTS or MTT) into a colored formazan product. These techniques are widely used to determine cell viability. MTS is cleaved in active mitochondria to a soluble formazan by mitochondrial succinate dehydrogenase which is located in complex II (succinate–ubiquinone oxidoreductase complex). The soluble formazan product is released to the culture medium; consequently, cultures can be assayed multiple times. 8.2.6.1 MTS Assay 1. Replace culture medium with phenol-free D-MEM. Let equilibrate for 30 min. 2. Add 20 µl of a solution of MTS and PMS (phenazine methosulfate), an electron coupler, as provided by the manufacturer, per 100 µl medium. 3. Return the culture to the incubator for 1 hr. 4. Record absorbance at 490 nm using an enzyne-linked immunosorbent assay (ELISA) plate reader. In the MTT assay, the formazan product is insoluble. For that reason, cell lysis and formazan solubilization are performed in an extra step. Maximum absorbance is read at 570 nm. Mitochondrial transmembrane potential was analyzed using JC1 (Molecular Probes, Eugene, OR).20,21 JC1 is a positively charged carbocyanine dye taken into the negatively charged inner mitochondrial membrane where it stays as a monomer at depolarized potential, producing a green fluorescence emission at 527 nm. In active mitochondria with higher membrane potentials, JC1 forms multimers called J aggregates that produce red fluorescence emissions at 590 nm. Hence, reduced mitochondrial activity results in reduced JC1 aggregation and decreased red fluorescence. Normal and DS astrocyte cultures were incubated with JC1 and analyzed by fluorescence microscopy. Cultures were treated with 0.25% trypsin for 5 min, dissociated and replated at a lower density (50,000 cells/cm2). After 24 hr, a dose–response curve was constructed to determine a nonsaturating concentration of JC1 in normal astrocyte cultures. This is important because at nonsaturating concentrations JC1 aggregation correlates with mitochondrial membrane potential, but at concentrations near or above saturation, the levels of J aggregate fluorescence rise linearly, independent of the mitochondrial membrane potential. Under our experimental conditions, a 30-min incubation with 0.5 µM JC1 resulted in approximately 70% of normal astrocytes exhibiting red fluorescent J aggregates. 8.2.6.2 JC1 Protocol 1. Incubate normal and DS astrocytes with 0.5 µM JC1 for 30 min. 2. Wash in warm PBS. 3. Quantify the number of individual astrocytes exhibiting J aggregates (emission at 590 nm) and the total number of astrocytes (emission at 527 nm) by image analysis of 10 microscopic fields per culture.

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Equivalent cellular loading of the dye in normal and DS cultures was confirmed by quantification of JC1 emission intensity at 527 nm. Equivalent astrocyte cell numbers in normal and DS cultures were further confirmed by scoring fluorescent nuclei after staining with Hoechst 33258. All experiments were performed blinded and in quadruplicate cultures. Approximately 450 cells were analyzed per experimental condition.

8.2.7 CELL VIABILITY ASSAYS Cell viability assays can be used during the setup of new cultures and to evaluate pharmacological treatments (neuroprotective or neurotoxic assays). Cell viability in cultures used for MTT, MTS, and JC1 assays was assessed using propidium iodide or trypan blue exclusion.7 These assays can be performed easily by microscopic analysis. 8.2.7.1 Trypan Blue Trypan blue is an acidic, water-soluble dye that cannot penetrate the intact plasma membranes of viable cells. If loss of membrane integrity occurs, trypan blue can enter cells and stain intracellular elements. Under bright field microscopic observation, live cells appear colorless and dead cells appear dark blue. Trypan blue is added to cells at 0.2% final concentration in D-MEM or serumfree media for 2 to 3 min. The culture is washed 3X with PBS, and the cells are observed under a bright field microscope at a final magnification of 100×. The number of stained cells is recorded in five to six fields per well in triplicate or quadruplicate wells. 8.2.7.2 Propidium Iodide Propidium iodide (PI) is a fluorescent dye that binds to double-stranded nucleic acids by intercalating between purines and pyrimidines, enhancing its fluorescent properties by 20- to 30-fold. It does not bind to single-stranded nucleic acids. The excitation and emission wavelengths of PI are ~493 and ~632 nm, respectively. Both excitation and emission are shifted ~35 and ~15 nm, respectively, after binding to nucleic acids. Similar to trypan blue, PI is cell-impermeable and can only penetrate inside a cell when the plasma membrane is damaged as a consequence of apoptotic or necrotic processes. After PI incubation, the culture is observed under fluorescent light using a 630-nm filter. The nuclei of nonviable cells appear red due to PI staining. Hoechst 33258, another dye that interacts with nucleic acids, can be used along with PI as a counterstain to assess the total number of cells in the culture. Hoechst has an excitation in the range of 351 to 363 nm and emission between 390 and 480 nm. PI is a known mutagen and care should be taken to use and dispose of it properly. Incubate cells with PI (10 µg/ml) in serum-free medium for 10 min. Wash cells gently with PBS. Fix the cells with 4% paraformaldehyde at 37oC for 20 min. Incubate fixed cells with Hoechst 33258 (10 µg/ml in PBS) for 5 min. Wash cells with PBS 3X. Mount the coverslip with antifade mounting media. Observe under

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fluorescence microscope with appropriate red filter for PI and UV filter for Hoechst 33258.

8.2.8 NEUROPROTECTION ASSAYS Treatment of DS neuronal cortical cultures growing in 24-well plates was initiated at day 7, before the onset of neuronal degeneration.7 A recombinant form of APP comprising amino acids 20 through 590 of APP695 and 17-mer reverse sequence peptides corresponding to a neurotrophic–neuroprotective domain of APP, kindly supplied by Dr. J.M. Roch22,23 (University of California, San Diego), were diluted in PBS and immediately added to the culture medium at the indicated concentrations. NGF, BDNF, and NT3 (supplied by Dr. M. Greenberg, Division of Neuroscience, Children’s Hospital, Boston, MA) were added to the culture medium at a final concentration of 40 ng/ml. At days 9 and 11, a 30% partial medium change was performed and the treatment was repeated. The cultures were fixed at day 14, immunostained with anti-α-tubulin class III, and the number of viable neurons was scored in quadruplicate cultures as previously described.7 More than 400 neurons were scored per experimental condition. For some experiments, DS neurons were incubated with conditioned medium of DS astrocyte cultures, normal astrocyte cultures or normal astrocyte-conditioned medium previously depleted of secreted APP by immunoprecipitation with a mixture of antibodies Alz-90 and 8E5.

8.3 RESULTS 8.3.1 DETECTION

OF INTRACELLULAR

β Aβ

IN

DS ASTROCYTES

To establish the presence and intracellular localization of Aβ, immunofluorescence microscopy was performed in DS astrocytes using monoclonal antibodies generated against the C termini of Aβ42 and Aβ40.17 These antibodies specifically recognized Aβ42 and Aβ40 respectively, but not full-length APP.16 Antibody specificity was also confirmed by immunocytochemical staining of synthetic Aβ1-42 and Aβ1-40 fibrils (data not shown). Intracellular Aβ42-specific staining was successfully achieved using a tyramide signal amplification system. Minimal background staining was detected in the cytoplasms of normal astrocytes (Figure 8.2A). In contrast, DS astrocytes had significant Aβ42-positive labeling in a vesicular distribution and often forming clusters in different regions of the cytoplasm (Figure 8.2B and Figure 8.2C, arrows). Aβ42 immunofluorescence was completely abolished by preincubation of the primary antibody with synthetic Aβ1-42 peptide (data not shown). Aβ42 was also detected in DS astrocytes extracted with Triton X-100 prior to fixation (Figure 8.2C), strongly suggesting the presence of insoluble, detergent-resistant intracellular Aβ42 aggregates. Aβ40 immunoreactivity was not detected in DS astrocytes. An electrophoretic gel system that separates Aβ40 from Aβ4224 confirmed the presence of intracellular Aβ42 in DS cortical cultures (data not shown).

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FIGURE 8.2 (See color insert following page 114.) Intracellular accumulation of aggregated Aβ42 in DS astrocytes. Immunofluorescence of normal (NL) and DS astrocyte cultures using α-Aβ42. Note the appearance of Aβ42 labeling in a vesicular-like distribution in DS (arrows; B and C) but not in NL astrocytes (A). Detergent-insoluble aggregates of Aβ42 appear in DS astrocytes extracted with Triton X-100 before fixation (C). Scale bar: 20 mm.

Western blotting and immunocytochemical analysis of Cu–Zn superoxide dismutase, another protein overexpressed in DS, and tubulin, an abundant cytoplasmic protein, showed no significant differences in protein levels and intracellular localization in DS cultures (data not shown), suggesting a specific mechanism in the

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accumulation and aggregation of intracellular Aβ (Aβi). Sequential double immunofluorescence showed that Aβi partially colocalized with APP, which is associated with subcellular compartments of the secretory pathway (Figure 8.3A). Aβi staining was particularly intense in perinuclear regions where it partially colocalized with the Golgi-resident protein syntaxin 6 (Syn), the early endosome antigen 1 protein (EEA1) and the lysosomal marker LAMP1 (Figure 8.3B through Figure 8.3D). Thus, intracellular Aβ42 accumulates in subcellular compartments associated with APP processing.

8.3.2 DS-LIKE ALTERATIONS IN APP PROCESSING INDUCED IN NORMAL ASTROCYTES BY ENERGY DEPLETION Alterations in APP metabolism observed in DS astrocytes16 are very similar to changes in APP processing induced by energy depletion.9 To further characterize the potential role of energy deficits in APP metabolism in the central nervous system (CNS), APP processing was analyzed in normal human astrocytes treated with the mitochondrial electron chain uncoupler CCCP.9,25 Energy depletion in normal astrocytes significantly increased the amount of cellular APP by 20.2 ± 4.3% (p < 0.002) and C99 by 50.6 ± 8.7% (p < 0.001) and reduced the levels of C83 and secreted APP (APPs) by 46.7 ± 7.7% (p < 0.001) and 49.6 ± 2% (p < 0.0001), respectively (Figure 8.4A and Figure 8.4B). Secreted Aβ levels were also reduced by CCCP treatment (data not shown). These changes are reminiscent of the alterations in APP processing in DS astrocytes. Immunocytochemical analysis of normal astrocytes after exposure to CCCP showed the presence of Aβ42 aggregates throughout the cytoplasm from perinuclear regions to the cell membrane (arrows, Figure 8.4C through Figure 8.4F). Aβ42 aggregates were resistant to solubilization with Triton X-100 (Figure 8.4F) and, similar to Aβ42 aggregates in DS astrocytes, partially co-localized with ER, Golgi and endosomal markers (data not shown). Thus, DS and energy-depleted normal astrocytes exhibit strikingly similar profiles of APP processing, including increased levels of cellular APP and C99, decreased levels of APPs and C83, and intracellular accumulation of detergent-resistant aggregates of Aβ42.

8.3.3 MITOCHONDRIAL DYSFUNCTION

IN

DS ASTROCYTES

The similarity in the alterations in APP processing observed in DS and CCCP-treated normal astrocytes raised the possibility that mitochondrial energy deficits could lead to altered APP processing in DS astrocytes. We investigated mitochondrial function by assessing mitochondrial transmembrane potential (∆Ψm) in viable DS and normal astrocytes using the fluorescent probe JC1.20,21 Normal and DS astrocyte cultures were incubated with JC1 and analyzed by fluorescence microscopy. Under these conditions, 72.9 ± 7.3% of normal astrocytes exhibited JC1 red fluorescence, indicative of active mitochondria (Figure 8.5A through Figure 8.5D). The number of DS astrocytes exhibiting J aggregates was significantly reduced to 42 ± 5.6% (Figure 8.5C and Figure 8.5D) despite similar cellular loading of JC1 as measured by JC1 emission intensity at 427 nm. Normal astrocytes treated with

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FIGURE 8.3 (See color insert following page 114.) Double labeling of Aβ42 and different subcellular markers. Double labeling shows partial colocalization of Aβ42 (Aβ) with APP (APP) in a DS astrocyte. APP immunostaining was performed with antibody Alz-90 (A). Double labeling shows partial colocalization of Aβ42 with the early endosomal antigen 1 (EEA1) in endosomal vesicles (B), with syntaxin 6 in the Golgi compartment (C), and with Lamp1 in lysosomal vesicles (D). Scale bar: 20 mm. (Panels A, B, and C reprinted from Busciglio, J. et al., 2002. With permission. Panel D prepared by the authors.)

CCCP showed a dramatic decrease in the number exhibiting J aggregates (16.7 ± 4.9%, Figure 8.5D). Thus, DS astrocytes exhibit significantly reduced mitochondrial transmembrane potential. To further examine mitochondrial function in DS, we assessed reduction of MTS reagent as an indicator of mitochondrial redox activity.26 MTS-reducing activity was

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FIGURE 8.4 (See color insert following page 114.) Mitochondrial dysfunction in normal astrocytes induces aberrant APP processing and intracellular Aβ accumulation. (A) Normal astrocytes treated with CCCP exhibit increased cellular APP (APP) and decreased levels of secreted APP (APPs). Samples were separated in a 4 to 20% gradient gel, electroblotted and incubated with antibodies C8 (cellular) and Alz-90 (secreted). (B) treatment with CCCP reduced C83 and increased C99 levels in normal astrocytes. Shown are Western blots with antibody C8 which recognizes APP C terminal fragments and 6E10 which recognizes C99 but not C83. (C through F) Accumulation of intracellular Aβ42 in CCCP-treated astrocytes. Immunofluorescence was performed on CCCP-treated and control astrocytes with antibody α-Aβ42. Nuclei were counterstained with propidium iodide. Intensely labeled aggregates of Aβ42 were localized throughout the cytoplasm after treatment with CCCP (arrows; D and F) and were detergent-insoluble (F). Scale bar: 20 mm. (Reprinted from Busciglio, J. et al., 2002. With permission.)

diminished by approximately 30% in DS cultures (p < 0.01, Figure 8.5E). A similar result was obtained using another mitochondrial redox assay utilizing the insoluble tetrazolium compound MTT (data not shown). Trypan blue and propidium iodide exclusion assays showed no significant differences in the number of viable cells in normal and DS cultures (Figure 8.5F), indicating that the decrease in mitochondrial redox activity in DS cultures was not due to decreased cell viability. Furthermore, analysis of cytochrome C levels by Western blotting did not show a significant difference between normal and DS astrocytes, suggesting that mitochondrial protein content was not different in DS cultures (data not shown). Taken together, these results suggest that mitochondrial energy metabolism is impaired in DS astrocytes.

8.3.4 APPS RESCUES DS CORTICAL NEURONS

FROM

APOPTOSIS

Previous studies suggest that APPs may generate both neurotrophic and neuroprotective activities.27 After the first week in culture,7 DS cortical neurons exhibited increased degeneration, accumulation of intracellular Aβ (Figure 8.2 and Figure 8.3) and decreased APP secretion.16 To determine whether APPs increase DS neuronal survival, cortical neurons were incubated with recombinant APP spanning residues 20 to 590 of APP69523 or a 17-mer peptide comprising the putative neurotrophic domain of APPs corresponding to amino acids 310 through 335.22,28 Incubation of DS cultures with either APP or the 17-mer peptide dramatically increased DS

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FIGURE 8.4 (continued)

neuronal survival (Figure 8.6A). In contrast, no significant effect was observed with a reverse sequence control peptide or with neurotrophic factors NGF, BDNF, and NT3 (Figure 8.6A). Thus, APPs (but not other neurotrophic factors) prevents the degeneration of DS neurons. APP did not significantly affect neuronal survival in normal cortical cultures (Figure 8.6A). To confirm a role for diminished APP secretion in DS neuronal degeneration, DS cortical neurons were incubated in culture with conditioned medium of normal or DS astrocytes from days 7 through 14. Normal astrocyte-conditioned medium dramatically increased DS neuronal survival, whereas DS astrocyte-conditioned medium did not increase survival (Figure 8.6B). DS cultures incubated with conditioned medium of normal astrocytes depleted of APP by immunoprecipitation did not exhibit increased neuronal survival (Figure 8.6B). Taken together, these results indicate that APPs acts as neuroprotectives agent for DS cortical neurons and that reduced levels of APPs may compromise neuronal survival in DS and AD.

8.4 DISCUSSION These experiments demonstrate intracellular Aβ accumulation in DS astrocytes that can be replicated in normal astrocytes by inhibition of mitochondrial energy metabolism. Moreover, mitochondrial function is impaired in DS astrocytes, as indicated

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FIGURE 8.5 (See color insert following page 114.) Impaired mitochondrial function in DS astrocytes. (A) Normal and DS astrocytes were labeled with the mitochondrial transmembrane potentialsensitive dye JC1 that fluoresces green (527 nm) or red (590 nm), depending on whether mitochondria are relatively less or more activated. A normal astrocyte (NL) labeled with JC1 shows inactive and active mitochondria labeled green and red, respectively, in the color insert. (B and C) DS astrocytes labeled with JC1 and visualized with the red channel show reduced numbers of cells with active mitochondria (arrows) and more cells with inactive mitochondria (arrowheads) relative to normal astrocytes. Astrocyte nuclei are labeled with Hoechst 33258 (blue channel in color insert). Scale bars: 20 mm. (D) Quantitative analysis shows a significant decrease in the number of DS astrocytes exhibiting red JC1 labeling (42 ± 5.6%) compared to normal astrocytes (72.9 ± 7.3%). CCCP-treatment of normal astrocytes markedly reduces the number of cells exhibiting red JC1 labeling (16.7 ± 4.9%). Values represent mean ± SEM; n = 4 independent experiments. *p < 0.01 relative to control by Student’s t test. (E) Reduced mitochondrial redox activity in DS astrocytes. MTS-reducing activity in DS cultures was significantly reduced relative to normal cultures. Values represent mean ± SEM; n = 4 independent experiments. *p < 0.01 relative to control by Student’s t test. (F) Trypan blue exclusion assay shows no significant differences in cell viability between normal and DS cultures. Values represent mean ± SEM; n = 4 independent experiments. (Panels B and C reprinted from Busciglio, J. et al., 2002. With permission. Panels A, D, E, and F prepared by the authors.)

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FIGURE 8.6 (A) Secreted APP prevents the degeneration of DS cortical neurons. DS cortical neuronal cultures were incubated in culture with recombinant APP (200 nM), an APP-derived trophic peptide (17-mer, 500 nm), the reverse sequence peptide (17-mer-rev, 500 nm) or neurotrophic factors NGF (40 ng/ml), BDNF (40 ng/ml) or NT3 (40 ng/ml) from days 7 to 14. Neuronal viability at day 14 is expressed as percent of the neuronal number at day 7 (100%). Neuronal viability in untreated DS cultures was reduced to 45 ± 4.8%. Recombinant APP and the 17-mer peptide prevented DS neuronal degeneration, whereas the 17-mer reverse peptide, NGF, BDNF, and NT3 did not affect DS neuronal survival. Values represent mean ± SEM; n = 4 independent experiments. *p < 0.01 by Student’s t test. (B) A significant increase in neuronal survival is observed in DS cultures incubated with conditioned medium of normal astrocyte cultures (c.m./NL APP(+)) but not with conditioned medium of DS astrocyte cultures (c.m./DS) or with conditioned medium of normal astrocytes depleted of APP by immunoprecipitation (c.m./NL APP(-)). Neuronal viability determined at day 14 in culture is expressed as percent of the neuronal number at day 7 (100%). Values represent mean ± SEM; n = 3 independent experiments. *p < 0.01 by Student’s t test.

by reduced mitochondrial redox activity and membrane potential. We suggest that impaired energy metabolism in DS cells gives rise to increased β-secretase cleavage of APP 16 and altered trafficking of Aβ, resulting in intracellular accumulation of aggregated Aβ42. Thus, impaired mitochondrial energy metabolism may contribute to AD pathogenesis in DS. We have also observed a similar impairment in mitochondrial function in primary cultures of DS fibroblasts (unpublished results), suggesting that mitochondrial dysfunction may be widespread in DS. We found intracellular Aβ in a detergent-resistant form in DS astrocytes and in normal astrocytes treated with the mitochondrial uncoupler CCCP. Intracellular Aβ42 partially colocalizes with APP and appears predominantly in the Golgi complex as

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well as in endosomal and lysosomal vesicles. It has previously been shown that intracellular Aβ42, APP and C-terminal fragments accumulate in a detergent-insoluble form in APP-overexpressing cells incubated with 20 µM synthetic Aβ1-42.29,30 Our results suggest that a similar phenomenon may occur under physiological conditions in DS astrocytes and in normal astrocytes under conditions of impaired energy metabolism. We have also detected detergent-insoluble APP that cosediments with intracellular Aβ in detergent-resistant pellets of DS cell lysates and brain homogenates (unpublished results). These experiments provide evidence for an alternative processing pathway in DS cells in which Aβ42, APP, and potentially amyloidogenic fragments accumulate and form detergent-resistant aggregates.16 A neuroprotective function of the secreted ectodomain of APP has been suggested by several studies.27,28,31 Our results show that APPs is significantly reduced in DS astrocytes in culture and in the DS brain.16 DS cortical neuronal cultures that exhibit reduced levels of APPs show a dramatic recovery in neuronal survival after incubation with recombinant APP or the 17-mer peptide corresponding to the neurotrophic domain of APP. DS neuronal survival was also significantly increased by incubation with conditioned medium of normal astrocytes, but not by the same medium depleted of APPs by immunoprecipitation. In contrast, NGF, BDNF, and NT3 did not improve DS neuronal survival. Taken together, these results suggest that APPs may be a survival factor for human neurons and underscore the potential pathological relevance of reduced APPs levels in DS and AD. In summary, we suggest that chronic APP overexpression may impair mitochondrial function, which in turn increases intracellular accumulation of Aβ and reduces secretion of neuroprotective APPs. Impaired mitochondrial function may result from a direct toxic effect of Aβ,32 APP accumulation in mitochondria,33 or from the metabolic cost of clearing aggregated proteins through chaperones and the degradative apparatus — processes that are highly energy-dependent. Future studies directed to analyze in detail the relationship between mitochondrial metabolism and neurodegeneration in DS are warranted.

ACKNOWLEDGMENTS This work was supported by grants from the National Institutes of Health (HD38466) and the Alzheimer’s Association to J.B. This chapter was reprinted partially from J. Busciglio et al., Altered metabolism of the amyloid beta precursor protein is associated with mitochondrial dysfunction in Down’s syndrome, Neuron, 33, 677, 2002. With permission from Elsevier.

REFERENCES 1. Coyle, J.T., Oster-Granite, M.L., and Gearhart, J.D. The neurobiologic consequences of Down syndrome, Brain Res. Bull., 16, 773, 1986. 2. Mann, D.M. The pathological association between Down syndrome and Alzheimer disease, Mech. Ageing Dev., 43, 99, 1988.

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3. Prasher, V.P. et al. Molecular mapping of Alzheimer-type dementia in Down’s syndrome, Ann. Neurol., 43, 380, 1998. 4. Selkoe, D.J. Alzheimer’s disease: genes, proteins, and therapy, Physiol. Rev., 81, 741, 2001. 5. Murphy, A.N., Fiskum, G., and Beal, M.F. Mitochondria in neurodegeneration: bioenergetic function in cell life and death, J. Cereb. Blood Flow Metab., 19, 231, 1999. 6. Beal, M.F. Aging, energy, and oxidative stress in neurodegenerative diseases, Ann. Neurol., 38, 357, 1995. 7. Busciglio, J. and Yankner, B.A. Apoptosis and increased generation of reactive oxygen species in Down’s syndrome neurons in vitro, Nature, 378, 776, 1995. 8. Zamzami, N. et al. Sequential reduction of mitochondrial transmembrane potential and generation of reactive oxygen species in early programmed cell death, J. Exp. Med., 182, 367, 1995. 9. Gabuzda, D. et al. Inhibition of energy metabolism alters the processing of amyloid precursor protein and induces a potentially amyloidogenic derivative, J. Biol. Chem., 269, 13623, 1994. 10. Blass, J.P. and Gibson, G.E. The role of oxidative abnormalities in the pathophysiology of Alzheimer’s disease, Rev. Neurol. (Paris), 147, 513, 1991. 11. Hirai, K. et al. Mitochondrial abnormalities in Alzheimer’s disease, J. Neurosci., 21, 3017, 2001. 12. Valla, J., Berndt, J.D., and Gonzalez-Lima, F. Energy hypometabolism in posterior cingulate cortex of Alzheimer’s patients: superficial laminar cytochrome oxidase associated with disease duration, J. Neurosci., 21, 4923, 2001. 13. Busciglio, J. et al. Generation of beta-amyloid in the secretory pathway in neuronal and nonneuronal cells, Proc. Natl. Acad. Sci. USA, 90, 2092, 1993. 14. Busciglio, J., Yeh, J., and Yankner, B.A. β-Amyloid neurotoxicity in human cortical culture is not mediated by excitotoxins, J. Neurochem., 61, 1565, 1993. 15. McConlogue, L. et al. Differential effects of a Rab6 mutant on secretory versus amyloidogenic processing of Alzheimer’s beta-amyloid precursor protein, J. Biol. Chem., 271, 1343, 1996. 16. Busciglio, J. et al. Altered metabolism of the amyloid beta precursor protein is associated with mitochondrial dysfunction in Down’s syndrome, Neuron, 33, 677, 2002. 17. Lippa, C.F. et al. Deposition of beta-amyloid subtypes 40 and 42 differentiates dementia with Lewy bodies from Alzheimer disease, Arch. Neurol., 56, 1111, 1999. 18. Pigino, G. et al. Presenilin-1 mutations reduce cytoskeletal association, deregulate neurite growth, and potentiate neuronal dystrophy and tau phosphorylation, J. Neurosci., 21, 834, 2001. 19. Heytler, P.G. Uncoupling of oxidative phosphorylation by carbonyl cyanide phenylhydrazones. I. Some characteristics of m-Cl-CCP action on mitochondria and chloroplasts, Biochemistry, 2, 357, 1963. 20. Reers, M. et al. Mitochondrial membrane potential monitored by JC-1 dye, Methods Enzymol., 260, 406, 1995. 21. Reers, M., Smith, T.W., and Chen, L.B. J-aggregate formation of a carbocyanine as a quantitative fluorescent indicator of membrane potential, Biochemistry, 30, 4480, 1991. 22. Roch, J.M. et al. Increase of synaptic density and memory retention by a peptide representing the trophic domain of the amyloid beta/A4 protein precursor, Proc. Natl. Acad. Sci. USA, 91, 7450, 1994. 23. Roch, J.M. et al. Bacterial expression, purification, and functional mapping of the amyloid beta/A4 protein precursor, J. Biol. Chem., 267, 2214, 1992.

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24. Wiltfang, J. et al. Improved electrophoretic separation and immunoblotting of betaamyloid (A beta) peptides 1-40, 1-42, and 1-43, Electrophoresis, 18, 527, 1997. 25. Fries, E. and Rothman, J.E. Transport of vesicular stomatitis virus glycoprotein in a cell-free extract, Proc. Natl. Acad. Sci. USA, 77, 3870, 1980. 26. Mosmann, T. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays, J. Immunol. Methods, 65, 55, 1983. 27. Mattson, M.P. Cellular actions of beta-amyloid precursor protein and its soluble and fibrillogenic derivatives, Physiol. Rev., 77, 1081, 1997. 28. Bowes, M.P. et al. Reduction of neurological damage by a peptide segment of the amyloid beta/A4 protein precursor in a rabbit spinal cord ischemia model, Exp. Neurol., 129, 112, 1994. 29. Yang, A.J. et al. Intracellular accumulation of insoluble, newly synthesized abetan42 in amyloid precursor protein-transfected cells that have been treated with Aβ1-42, J. Biol. Chem., 274, 20650, 1999. 30. Yang, A.J. et al. Intracellular A beta 1-42 aggregates stimulate the accumulation of stable, insoluble amyloidogenic fragments of the amyloid precursor protein in transfected cells, J. Biol. Chem., 270, 14786, 1995. 31. Mattson, M.P. et al. Evidence for excitoprotective and intraneuronal calcium-regulating roles for secreted forms of the beta-amyloid precursor protein, Neuron, 10, 243, 1993. 32. Casley, C.S. et al. Beta-amyloid inhibits integrated mitochondrial respiration and key enzyme activities, J. Neurochem., 80, 91, 2002. 33. Anandatheerthavarada, H.K. et al. Mitochondrial targeting and a novel transmembrane arrest of Alzheimer’s amyloid precursor protein impairs mitochondrial function in neuronal cells, J. Cell Biol., 161, 41, 2003.

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9

Linking Alzheimer’s Disease, β-Amyloid, and Lipids: A Technical Approach Marcus O.W. Grimm, Andreas J. Paetzold, Heike S. Grimm, Eva G. Zinser, Thomas Ruppert, and Tobias Hartmann

CONTENTS 9.1 9.2

9.3

9.4

Introduction Cholesterol and Aβ . 9.2.1 Cell Culture and Cholesterol Depletion 9.2.2 Protein Analysis 9.2.3 Results Sphingolipids and AD 9.3.1 Cell Culture 9.3.2 Enzymatic Assay 9.3.3 Lipid Extraction 9.3.4 Phosphorus Determination 9.3.5 Scintillation Count Analysis of Lipids by Mass Spectrometry 9.4.1 Theoretical Background 9.4.2 Sample Preparation 9.4.3 Sample Application 9.4.4 Measurement 9.4.4.1 Instrumentation 9.4.4.2 TOF pos Measurement . 9.4.4.3 Precursor Ion Scans on m/z = 184.1, 188.1, and 264.3 9.4.5 Data Interpretation 9.4.6 Key Steps and Modifications 9.4.6.1 Sample Preparation 9.4.6.2 Sample Application

0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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9.4.6.3 TOF pos Measurement 9.4.6.4 Precursor Ion Scan 9.4.7 Results Acknowledgment References

9.1 INTRODUCTION Until recently, four genes harboring point mutations that significantly affect Alzheimer’s disease (AD) pathogenesis have been identified.1 All these mutations result in increased β-Amyloid 42 (Aβ42) levels.2 Three of these genes, the amyloid precursor protein (APP), presenilin 1 (PS1), and presenilin 2 (PS2), are involved in the molecular pathway of AD. The fourth gene, apolipoprotein E (ApoE), suggests a possible link to lipid pathways. ApoE encodes a lipid-binding protein that transports lipids between cells and is therefore an important factor in lipid homeostasis.3 Human ApoE exists in three major alleles. In epidemiological studies, the ε4 allele increases the risk for hypercholesterolemia and decreases the age of onset of AD.4 Moreover, ApoE transgenic and knockout mice show altered Aβ deposition indicating that ApoE and lipids may play a significant role in Aβ pathology.5 Alterations in lipid homeostasis have long been recognized to severely affect neuronal function and cause neurodegenerative diseases.6 Table 9.1 presents a summary of the most common lipid-related diseases. Enzymes of the sphingolipid and ganglioside pathways seem to be involved in neurodegenerative disorders. e.g., in Niemann-Pick disease, the sphingomyelin level is drastically increased by a lack of the sphingomyelin-degrading enzyme, the acid sphingomyelinase (Figure 9.1). However, it should be noted that these diseases are often fatal during childhood or early adulthood and therefore little is known about their relevance to AD. APP and all APP secretases are transmembrane proteins. Therefore it is conceivable that lipids may also play a fundamental role in Alzheimer’s disease. It has recently been shown that intramembrane proteolysis of APP is influenced by the physical characteristics of the membrane. For example the γ-cleavage seems to take place in the middle of the transmembrane domain of APP.7 Aβ was still generated when residues within the transmembrane domain were mutated,8 whereas the cleavage site shifted by an altered length of the transmembrane domain, which led to a changed ratio of Aβ42 to Aβ40. C terminal insertion of two residues and N-terminal deletion of two residues strongly altered the ratio. As the length of the transmembrane domain corresponds to the membrane diameter, it is obvious that a changed membrane thickness may produce drastic consequences for the Aβ42/Aβ40 ratio.8 Accordingly, a membrane that has a smaller diameter should shift toward Aβ42 production, which is in fact the case for comparisons of plasma and endoplasmic reticulum (ER) membranes. In summary, this model suggests that the ratio of pathogenic Aβ42 and nonpathogenic Aβ40 is modulated by membrane composition.

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TABLE 9.1 Lipid-Related Neuronal Disorders and Affected Lipids and Enzymes Disease Faber disease Niemann-Pick disease Krabbe disease

Affected Lipids

GM1-gangliosidosis

Ceramide Sphingomyelin Galactosylceramide, Galactosylsphingosine Glucosylceramide, Glucosylsphingosine Digalactosylceramide GM-ganglioside GM-ganglioside Sulfatide Sulfatide Sulfatide, globotriaosylceramide, digalactosylceramide, GM3-ganglioside Glucosylceramide All glycolipids with short sugarchains, e.g., Cer, GlcCer, LacCer, GalCer, DigalCer, Sulfatide GM1-ganglioside

GM2-gangliosidosis (B1 variant) GM2-gangliosidosis (AB variant)

GM2-ganglioside GM2-ganglioside

Gaucher disease Fabry disease Tay-Sachs disease Sandhoff disease Metachoromatic leukodystrophy Multiple sulfatase deficiency Sulfatidase activator deficiency (sap-B deficiency) SAP-2 deficiency SAP precursor deficiency

Enzymatic Defects Acid ceramidase Sphingomyelinase Galactosylceramidase Glucosylceramidase α-Galactosidase A β-Hexosaminidase A β-Hexosaminidases A and B Arylsulfatase A (sulfatidase) Arylsulfatases A, B, and C Sulfatidase activator (SAP-1, sap-B) SAP-2 (sap-C) SAP precursor, sap-A, B, C, and D

GM1-ganglioside, β-Galactosidase β-Hexosaminidase A β-Hexosaminidase A

9.2 CHOLESTEROL AND Aβ Animals fed a cholesterol-enriched diet showed increased accumulation of Aβ in their brains.9 Similarly, reduced cellular cholesterol levels resulted in decreased Aβ production.10 Primary neurons treated with cholesterol lowering drugs, e.g., statins, produced significantly reduced Aβ levels.11 In order to evaluate whether unspecific side effects of the statins or indeed cholesterol depletion caused reduced Aβ production, cholesterol plasma membrane levels were decreased with methyl-β-cyclodextrin (CDX).12 While statins reduced cholesterol levels by inhibition of 3-hydroxy-3methyl-glutaryl coenzyme A (HMG-CoA) reductase, CDX physically extracted cholesterol from the plasma membranes. These mechanistically unrelated approaches led to similarly decreased Aβ levels, indicating that cholesterol levels and not a side effect of the statins are responsible for altered Aβ production.13 Treatment of guinea pigs with a high dosage of Simvastatin induced a reversible decrease in cerebrospinal fluid and brain tissue Aβ levels.14 Similar results were found using the cholesterol synthesis inhibitor BM15.766 which inhibits ∆7-reductase, a later step in cholesterol synthesis15,16 (Figure 9.2).

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FIGURE 9.1 Biochemical pathway of sphingomyelin synthesis. The names of the compounds appear in large type; enzyme names appear in smaller type.

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FIGURE 9.2 Biochemical pathway of cholesterol synthesis. The names of the compounds appear in bold type; enzyme names appear in normal type. Inhibitors are noted at places of action. BM15.766 is blocking the ∆7-reductase that catalyzes one of the final steps of the pathway.

© 2005 by Taylor & Francis Group.

The following sections will describe the experimental approaches to lowering cholesterol levels in cell cultures.

9.2.1 CELL CULTURE

AND

CHOLESTEROL DEPLETION

As mentioned above, cholesterol level can be decreased by inhibition of HMG-CoA reductase as a committed step enzyme in cholesterol biosynthesis or by physical depletion of plasma membrane cholesterol. To maximize the effect on Aβ production, these approaches can be used in combination. Hippocampal neurons were prepared from 17-day-old fetal rats.17 The neurons were cultivated for 5 days at 90% humidity and 5% CO2 atmosphere at 37˚C in N2MEM. Subsequently 4 µM lovastatin (Calbiochem, La Jolla, CA) or simvastatin (Merck, Whitehouse Station, NJ) and 0.25 mM mevalonate (Sigma Chemical, St. Louis, MO) were added for 48 or 72 hr. By supplementing the medium with 5 mM mevalonate, biochemical pathways for nonsteroidal products were allowed to proceed. Without this supplementation cellular toxicity increased.14,18 The neurons were infected with SFV-APP695 for 60 to 90 min, depending on the virus titer. Exposition times longer than 90 min are not recommended because of increasing cytotoxicity. After washing the cells with N2MEM and incubation for 2 hr, the cells were treated with 5 mM CDX (Sigma) for 10 min. With longer exposure times or higher concentrations of CDX, Aβ levels dropped below the detection limit. After 3.5 to 4.5 hr inside the nontoxic window of SFV infection, cells were harvested and Aβ levels were analyzed. As a control, cells were treated in the same way in the absence of CDX and statins. To analyze the effects of statins without cyclodextrin, cyclodextrin treatment was omitted as a variation of the experiment. Moreover, to avoid analyzing effects limited to hippocampal neurons, mixed cortical neurons from brain were used.

9.2.2 PROTEIN ANALYSIS Cell culture media were collected and cell extracts were prepared in 2% (v/v) Nonidet P-40, 0.2% (w/v) SDS, and 5 mM EDTA supplemented with complete protease inhibitor (Roche, Basel, Switzerland). In order to detect equal intensities of all analyzed Aβ-forms, cell lysates were divided differently for Aβ42 (79%) and Aβ40 (19%). The remaining 2% was saved for direct loading of cell lysates for detection of APP and total Aβ. The conditioned media were split 90% for Aβ42 and 10% for Aβ40 production. Adjusting Aβ levels to equal ranges of band intensities for Aβ40 and Aβ42 allows a more precise determination of Aβ ratio because densiometric analysis of the Western blot can be performed from individual blots and exposition times. For immunoprecipitation, monoclonal antibodies G2-10 (2 µg/ml) for Aβ40 and G2-11 (4 µg/ml) for Aβ42 were used. The monoclonal W0-2 antibody directed against amino acids 4 through 10 of human Aβ was used for total Aβ and APP detection. Quantitative immunoprecipitation was performed according to Schröder et al.19 Precipitates were analyzed on 10 to 20% Tris-Tricine polyacrylamide gels (Novex,

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San Diego, CA). Quantitative Western blotting was performed modified to Ida et al.29 For blocking 3% (w/v) milk powder instead of BSA was used. G2-10 and G2-11 precipitates were detected with W0-2 antibody. Quantification was done after enhanced chemoluminescence (ECL) development by densiometric scans.

9.2.3 RESULTS De novo synthesis of cholesterol was blocked by both lovastatin and simvastatin. Additionally, plasma membrane cholesterol was extracted by CDX. In the conditioned media, cholesterol depletion by lovastin and CDX decreased Aβ40 levels to 50% (±13%). Aβ42 levels were decreased to 29% (±10%) compared to untreated cells. Similar results were obtained for intracellular Aβ levels. Aβ40 levels were decreased to 42% (±18%) and intracellular Aβ42 levels dropped to 33% (±15%) compared to untreated cells. A more detailed description of the results can be found in Fassbender et al.14

9.3 SPHINGOLIPIDS AND AD As described above, sphingolipids and gangliosides play important roles in many neurodegenerative disorders. Sphingolipids and cholesterol play crucial roles in lipid raft biology and modulate the physical states of membranes.20,21 Moreover, it has been recently found that sphingolipid levels and composition are altered in AD brains, indicating a role for this lipid class in AD pathology.22 The following section describes experiments analyzing the committed step reaction of the sphingolipid pathway catalyzed by serine–palmitoyl–transferase (SPT). This enzymatic reaction catalyzes the condensation of serine and palmitoyl-CoA, a reaction that produces 3-ketodihydrosphingosine (KDS). SPT belongs to a family of pyridoxal 5′-phosphate (PLP)-dependent α-oxoamine synthases (POAS). Mammalian SPT is a heterodimer of 53-kDa LCB1 and 63-kDa LCB2 subunits localized at the ER. The enzymatic assay is based on radioactive incorporation of 14C-serine. While 14C-serine is water soluble, the originated products of SPT and the enzymes of the sphingolipid pathway are liposoluble. As a control, the same approach is performed with 1 mM myriocin. Myriocin is a selective inhibitor of the SPT. Under these conditions no activity should occur.

9.3.1 CELL CULTURE Cells were cultured in DMEM containing 5 to 10% fetal calf serum (FCS) at 37˚C, 90% humidity, and 5% CO2 atmosphere. Cells were grown until a 90% confluent cell layer appeared.

9.3.2 ENZYMATIC ASSAY Cells were washed three times with Buffer 1 containing 100 mM HEPES, pH 7.3. After harvesting, the cells were homogenized in 100 mM HEPES, pH 7.3, supplemented with complete protease inhibitor (Roche). Homogenization and the following

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steps were performed at 4˚C in order to decrease protein degradation. The homogenates were adjusted to equal protein levels. For each assay, a protein concentration of 5 to 7 mg/ml in a final volume of 3000 µl is recommended. Lower protein concentrations result in enzymatic activity that is too low for proper analysis. The reaction is started by adding 300 µl of Buffer 2 containing 500 µM pyridoxalphosphate (Sigma), 100 mM HEPES, pH 7.3, 3 mM palmitoyl-CoA, and 10 µCi/ml L-serine (Amersham Biosciences, Uppsala, Sweden) at 37˚C in glass tubes (Wheaton, Millville, NJ). To stop the reaction 400 µl of the reaction mixture is transferred in glass tubes containing 3.75 ml freshly prepared solvent CHCl3:MeOH:HCl (5:10:0.075). The reaction is stopped after 0, 2, 4, 8, 16, 32, 64, and 96 min. It is very important to use freshly prepared solvent because the very low vapor pressure of chloroform leads to a high rate of evaporation of chloroform and changed ratios of the different solvents. Glass tubes instead of plastic ones must be used because lipids adhere to plastic surfaces.

9.3.3 LIPID EXTRACTION The lipid extraction was based on the method of Bligh and Dyer23 with some modifications that are briefly described. All steps were performed at room temperature. The samples were vortexed in the glass tubes for 60 min. After 1 ml CHCl3 was added, the samples were vortexed again for 10 min. Phase separation occurred by the addition of 1 ml distilled H2O. The resulting hydrophilic and lipophilic phases were thoroughly mixed by vortexing for 30 min, after which the phases were separated by centrifugation for 10 min at 3000 × g. The lower lipophilic phase was transferred to a new glass tube. The transfer of the lower lipophilic phase must be performed quantitatively without contaminating the organic phase with the interphase or hydrophilic phase. The interphase and the hydrophilic phase contain nucleic acids, carbohydrates, and proteins, whereas the lipophilic phase contains lipids, especially sphingolipids and phospholipids. The procedure is carried out two more times starting again with the addition of CHCl3:MeOH:HCl (5:10:0.075). After the last extraction, the lipophilic phase containing the extracted lipids, is evaporated under continuous nitrogen flow. The samples were resolved again in 200 µl CHCl3:MeOH:HCl (5:10:0.075) by vortexing for 60 min; 50 µl are needed for phosphorus determination and the remaining 150 are applied for scintillation count.

9.3.4 PHOSPHORUS DETERMINATION It should be noted that phosphate-based detergents must not be used for glassware when this method is employed. All glassware should be washed with distilled H2O immediately before use. As noted earlier, the lipophilic phase contains most of the lipids and especially phospholipids. The phospholipid concentrations of the different samples should not differ after extraction.

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Analyzing phosphorus concentration that directly corresponds to phospholipid concentration reveals the quality of the extraction and points out errors due to extraction. In order to quantify the inorganic phosphorus concentration, the phospholipids were digested by refluxing in perchloric acid to release inorganic phosphate. The inorganic phosphate was then converted to phosphomolybdic acid which was reduced to a blue compound for spectrometric determination.24 To prepare the chromogenic solution, 16 g ammonium molybdate is dissolved in 120 ml water (Reagent A), after which 40 ml of hydrochloric acid and 10 ml mercury are added to 80 ml of Reagent A and thoroughly mixed. The supernatant is used as Reagent B; 40 ml of Reagent A is combined with 200 ml concentrated sulfuric acid and added to Reagent B to produce Reagent C. Subsequently, 25 ml of Reagent C is added to 45 ml methanol, 5 ml chloroform, and 20 ml water. This chromogenic solution can be stored at 4˚C for several weeks. The next step is adding 13 µl chromogenic solution to 50 µl sample and the solution is heated at 100˚C for 75 sec. After the samples are thoroughly mixed and cooled for 5 min to room temperature, 500 ml nonane is added. The samples are briefly mixed and incubated for 15 min. The tubes are centrifuged for 3 min at 3000 × g before the absorbance of the supernatant at 730 nm is compared with the blank where 50 µl CHCl3:MeOH:HCl (5:10:0.075) is used instead of the sample. Very high amounts of cholesterol influence the phosphorus determination.

9.3.5 SCINTILLATION COUNT The remaining 150 µl of the sample is used for scintillation counting. First, 2.5 ml of scintillation mix is added and thoroughly mixed for 1 hr, after which its radioactivity is determined in a scintillation counter (Beckmann LS 6000IC). Figure 9.3 shows serine–palmitoyl–transferase (SPT) activity in murine fibroblast cells.

FIGURE 9.3 Serine-palmitoyl-transferase (SPT) activity in murine fibroblast cells.

© 2005 by Taylor & Francis Group.

9.4 ANALYSIS OF LIPIDS BY MASS SPECTROMETRY Alzheimer’s disease is linked to lipid homeostasis. Therefore this is a reliable method to identify and quantify sphingomyelin and ceramide. In recent years, mass spectrometry has become a well-established biochemical method, particularly after the genomic research led to the use of mass spectrometry to identify different proteins. The subsequent sections will focus on the usage of mass spectrometry in lipid analysis.

9.4.1 THEORETICAL BACKGROUND Electrospray ionization mass spectrometry (ESI-MS) is a very efficient tool for the analysis of lipid extracts.25–28 Lipids dissolved in an appropriate solvent are introduced through a small-diameter needle to which a high voltage is applied. At the needle tip, the surface of the solution becomes highly charged and droplets are formed to increase the surface. The droplets shrink due to evaporation of solvent which increases the charge density at the surface. When the surface charge density exceeds the Rayleigh stability limit, electrostatic repulsion greater than the surface tension of the droplet results in an explosion of the droplet and expulsion of gas phase ions. These ions are positively or negatively charged, depending on the polarity of the applied voltage. In the following method, lipids are characterized by a quadrupole time-of-flight mass analyzer (Q-TOF) consisting of a quadrupole that serves as a mass filter, a collision cell for collision-induced fragmentation and a TOF analyzer. Mass determination of positive charged lipid ions is performed using only the TOF analyzer (TOF pos). Ions are accelerated in an electric field and the time is determined until the ions reach the detector. From the time of flight, the mass-to-charge ratio (m/z value) of the ions is calculated. Structural information of a particular lipid ion is obtained by selecting the ions using the first quadrupole. Only these ions enter the collision cell. After collision with nitrogen molecules, the generated fragment masses are determined by the TOF analyzer (product ion scan). Using the first quadrupole in a scanning mode, ions of a complex mixture are allowed to enter the collision cell sequentially, depending on their mass. After fragmentation, the TOF analyzer is used to detect the appearance of a characteristic fragment and the masses of components are calculated from which this fragment is derived (precursor ion scan or PIS). Fragmentation of sphingomyelins yields a fragment of m/z = 184.1 that is characteristic for the phosphorylcholine moiety (see Figure 9.4 and References 25 and 26). This fragment is not characteristic for sphingomyelins because it also results from fragmentation of any representative of phosphatidylcholines (PCs). However, although monoprotonated sphingomyelins have odd nominal m/z values, phosphatidylcholines have nominal m/z values because of a formal exchange of an amide group (15 Da) for an oxygen atom (16 Da). Furthermore, fragmentation of sphingomyelin and of ceramides leads to a fragment of m/z = 264.3 (see Figure 9.4), representing the di-dehydro-sphingenine body. This is characteristic for ceramides and is built up by the fusion of serine and palmitoyl-CoA in the sphingolipid biosynthesis pathway (compare Figure 9.1). By comparison of a simultaneous PIS

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FIGURE 9.4 (A) TOF spectrum of a positive mode fragmentation of N-palmitoyl-D-erythro-sphingosyl-phosphorylcholine (sphingomyelin 16:0) in the mass range from m/z = 70 to 720. The mass range which is extended 10 times (does not include the peak at 184.07) is labeled ×10,0 above the spectrum. Each peak is labeled with its centroid mass. (B) Compounds with corresponding masses including their paths of formation during fragmentation of sphingomyelin 16:0.

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on m/z = 184.1 and 264.3 in the presence of an external standard, it is possible to determine and quantify sphingomyelins and ceramides.

9.4.2 SAMPLE PREPARATION The procedure of sample preparation is the same as described above. However, the recommendation is to use 25 to 50% of the cells as in the case of thin layer chromatography. For mass spectrometry, the solvent should be CHCl3:MeOH:HCl (5:10:0.075). If necessary evaporate a different solvent under nitrogen atmosphere and resolve the sample in CHCl3:MeOH:HCl (5:10:0.075). The addition of approximately 1.8 pmol 1,2-dimyristoyl-sn-glycero-3-phosphocholine-1,1,2,2-D4 (Avanti Polar Lipids, Inc., Alabaster, AL) as an external standard will produce a reference peak of m/z = 682.5 in TOF pos spectra.

9.4.3 SAMPLE APPLICATION 1. Rinse a shortened nano-ESI needle (Econo12, New Objective, Woburn, MA) with approx. 5 µl of sample. 2. Subsequently fill the needle with 5 to 10 µl of sample. 3. Feed it into the holder of the ESI source. 4. The needle tip should be placed about 1 mm from the MS orifice.

9.4.4 MEASUREMENT 9.4.4.1 Instrumentation Spectrometric analysis was performed on a Q-TOF Pulsar mass spectrometer (PE SIEX, Weiterstadt, Germany) equipped with a nano-ESI source (MDS Protana, Odense, Denmark). 9.4.4.2 TOF pos Measurement 1. 2. 3. 4.

Charge nano-ESI needle with a potential of 700 V. Set declustering potential to 40 eV. Set TOF pos range to m/z = 600 to 900. While monitoring the spectrum, note whether total ion count (TIC) correlates with extracted ion count (XIC) of reference peak (m/z = 682.5) and at least one peak representing sphingomyelin, e.g., m/z = 703.3 or 731.3. 5. Maintain a stable spectrum for at least 2 min. 9.4.4.3 Precursor Ion Scans on m/z = 184.1, 188.1, and 264.3 1. 2. 3. 4.

Set TOF range to m/z = 170 to 270. Enable peak enhancement for m/z = 264.3. Set collision energy to 60 eV. Run PIS in ranges of m/z = 500 to 690 and 670 to 820, respectively.

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9.4.5 DATA INTERPRETATION TOF pos spectra were calculated by Analyst QS software (Applied Biosystems, Foster City, CA) by averaging a 2-min period of the profile. Precursor ion scans were calculated in the same way for at least 5 min. Further interpretations were made after exporting Analyst QS data into MS Access. A database was programmed to pair same peaks from different spectra with a ∆m/z of 0.05 and normalize intensities to the reference peak of the added external standard m/z = 682.5 or 188.1, respectively.

9.4.6 KEY STEPS

AND

MODIFICATIONS

9.4.6.1 Sample Preparation The reduced number of cells is recommended to reduce the possibility of formation of precipitates on the tip of nano-ESI needle. Our dilution experiments indicate that these precipitates are not based on high concentrations of lipids or salts. 9.4.6.2 Sample Application Rinsing of the needle is recommended to achieve a stable spray in a shorter time. 9.4.6.3 TOF pos Measurement It is essential for quantification that the spray is stable, i.e., the spectrum does not change in the course of testing. Additionally, the precursor ion scan needs a reasonable signal input. 9.4.6.4 Precursor Ion Scan The characteristic fragment ions of phosphorylcholines, m/z = 184.1, and ceramide, m/z = 264.3, were selected for discrimination of phosphatidylcholines from ceramides and/or sphingomyelins.

9.4.7 RESULTS Fragmentation of the external standard mentioned above leads to a peak at m/z = 188.1 instead of m/z = 184.1. A precursor ion scan on m/z = 188.1 results in a maternal peak of m/z = 682.5 and its first and second isotope peaks (Figure 9.5). This clear-cut result shows the suitability of the external standard and the conditions.

ACKNOWLEDGMENTS We would like to thank Inge Tomic for technical support and Marco Duering for helpful advice.

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FIGURE 9.5 Partial screen shot from Analyst QS software. It is an example of a precursor ion scan (PIS) approach to analyzing a lipid extract derived from a cell homogenate of a retransfected preseniline double knockout cell line. (A) Total ion counts per time (TIC) representing the stability of the spray. (B) PIS of the fragment m/z = 184.1 (phosphatidylcholine headgroup). (C) Result of the simultaneous PIS of the head group (m/z = 188.1) of standard (1,2-dimyristoyl-sn-glycero-3phosphocholine-1,1,2,2-D4). (D) Peaks generated by PIS of di-dehydro-sphingenine (m/z = 264.3).

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REFERENCES 1. De Strooper, B. and Annaert, W. Proteolytic processing and cell biological functions of the amyloid precursor protein. J. Cell Sci. 113, 1857–1870, 2000. 2. Sinha, S. and Lieberburg, I. Cellular mechanisms of beta-amyloid production and secretion. Proc. Natl. Acad. Sci. USA 96, 11049–11053, 1999. 3. Mahley, R.W. and Rall, S.C., Jr. Apolipoprotein E: far more than a lipid transport protein. Annu. Rev. Genomics Hum. Genet. 1, 507–337, 2000. 4. Corder, E.H. et al. Gene dose of apolipoprotein E type 4 allele and the risk of Alzheimer’s disease in late onset families. Science 261, 921–923, 1993. 5. Bales, K.R. et al. Lack of apolipoprotein E dramatically reduces amyloid beta-peptide deposition. Nat. Genet. 17, 263–264, 1997. 6. Nathan, B.P. et al. Differential effects of apolipoproteins E3 and E4 on neuronal growth in vitro. Science 264, 850–852, 1994. 7. Grziwa, B. et al. The transmembrane domain of the amyloid precursor protein in microsomal membranes is on both sides shorter than predicted. J. Biol. Chem. 278, 6803–6308, 2003. 8. Lichtenthaler, S.F. et al. The intramembrane cleavage site of the amyloid precursor protein depends on the length of its transmembrane domain. Proc. Natl. Acad. Sci. USA 99, 1365–1370, 2002. 9. Sparks, D.L. et al. Induction of Alzheimer-like beta-amyloid immunoreactivity in the brains of rabbits with dietary cholesterol. Exp. Neurol. 126, 88–94, 1994. 10. Simons, M. et al. Cholesterol depletion inhibits the generation of beta-amyloid in hippocampal neurons. Proc. Natl. Acad. Sci. USA 95, 6460–6464, 1998. 11. Cucchiara, B. and Kasner, S.E. Use of statins in CNS disorders. J. Neurol. Sci. 187, 81–89, 2001. 12. Kilsdonk, E.P. et al. Cellular cholesterol efflux mediated by cyclodextrins. J. Biol. Chem. 270, 17250–17256, 1995. 13. Frears, E.R. et al. The role of cholesterol in the biosynthesis of beta-amyloid. Neuroreport 10, 1699–1705, 1999. 14. Fassbender, K. et al. Simvastatin strongly reduces levels of Alzheimer’s disease betaamyloid peptides Abeta 42 and Abeta 40 in vitro and in vivo. Proc. Natl. Acad. Sci. USA 98, 5856–5961, 2001. 15. Refolo, L.M. et al. Hypercholesterolemia accelerates the Alzheimer’s amyloid pathology in a transgenic mouse model. Neurobiol. Dis. 7, 321–3331, 2000. 16. Refolo, L.M. et al. A cholesterol-lowering drug reduces beta-amyloid pathology in a transgenic mouse model of Alzheimer’s disease. Neurobiol. Dis. 8, 890–899, 2001. 17. Tienari, P.J. et al. Intracellular and secreted Alzheimer beta-amyloid species are generated by distinct mechanisms in cultured hippocampal neurons. Proc. Natl. Acad. Sci. USA 94, 4125–4130, 1997. 18. Fassbender, K. et al. Effects of statins on human cerebral cholesterol metabolism and secretion of Alzheimer amyloid peptide. Neurology 59, 1257–1258, 2002. 19. Schroder, J. et al. Cerebral changes and cerebrospinal fluid beta-amyloid in Alzheimer’s disease: a study with quantitative magnetic resonance imaging. Mol. Psychiatr. 2, 505–507, 1997. 20. Roy, D. and Mukhopadhyay, C. GD1a in phospholipid bilayer: a molecular dynamics simulation. J. Biomol. Struct. Dyn. 18, 639–646, 2001. 21. Kappel, T. et al. Gangliosides affect membrane-channel activities dependent on ambient temperature. Cell. Mol. Neurobiol. 20, 579–590, 2000.

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22. Cutler, R.G. et al. Involvement of oxidative stress-induced abnormalities in ceramide and cholesterol metabolism in brain aging and Alzheimer’s disease. Proc. Natl. Acad. Sci. USA 101, 2070–2075, 2004. 23. Bligh, E.G and Dyer, W.J. A rapid method of total lipid extraction and purification. Can. J. Med. Sci. 37, 911–917, 1959. 24. Hundrieser, K.E., Clark, R.M., and Jensen, R.G. Total phospholipid analysis in human milk without acid digestion. Am. J. Clin. Nutr. 41, 988–993, 1985. 25. Brugger, B. et al. Quantitative analysis of biological membrane lipids at the low picomole level by nano-electrospray ionization tandem mass spectrometry. Proc. Natl. Acad. Sci. USA 94, 2339–2344, 1997. 26. Ekroos, K. et al. Charting molecular composition of phosphatidylcholines by fatty acid scanning and ion trap MS3 fragmentation. J. Lipid Res. 44, 2181–2192, 2003. 27. Han, X. and Gross, R.W. Electrospray ionization mass spectroscopic analysis of human erythrocyte plasma membrane phospholipids. Proc. Natl. Acad. Sci. USA 91, 10635–10639, 1994. 28. Kerwin, J.L., Tuininga, A.R., and Ericsson, L.H. Identification of molecular species of glycerophospholipids and sphingomyelin using electrospray mass spectrometry. J. Lipid Res. 35, 1102–1114, 1994. 29. Ida, N. et al. Analysis of heterogeneous A4 peptides in human cerebrospinal fluid and blood by a newly developed sensitive Western blot assay. J. Biol. Chem. 271, 22908–22914, 1996.

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10

Regulation of Amyloid Precursor Protein Processing by Lithium Xiaoyan Sun and Akihiko Takashima

CONTENTS Abstract 10.1 Introduction 10.2 Experimental Procedures 10.2.1 Lithium Treatment in Transfected Cultured Cells 10.2.2 Aβ ELISA 10.2.3 Preparation of fibrillar Aβ (fAβ) 10.2.4 Microinjection of fAβ into Tau Transgenic Mice 10.2.5 Lithium Administration 10.3 Results 10.3.1 Establishment of Selective Sandwich Aβ ELISA 10.3.2 Effect of LiCl on Aβ Secretion in Association with GSK-3β Activity from APP C100-Transfected COS7 Cells 10.3.3 Effect of GSK-3β on fAβ-Induced Tau Pathology In Vivo 10.4 Discussion References

ABSTRACT Secreted Aβ is produced during normal cell metabolism and it is regulated by many factors. Quantification of Aβ generation is an important approach to studying APP processing. In this chapter, we demonstrate that lithium, a potent GSK-3 inhibitor, can regulate amyloid secretion in COS7 cells overexpressing APPC100 demonstrated by a sandwich enzyme-linked immunosorbent assay (ELISA). Finally, we show that lithium can effectively inhibit tau pathology induced by Aβ injection into the hippocampus of a transgenic mouse model.

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10.1 INTRODUCTION Alzheimer’s disease (AD) is characterized by the deposition of amyloid-β peptide (Aβ) and accumulation of hyperphosphorylated tau protein in patients’ brains. Accumulating evidence indicates that several kinases play an important role in the development of AD pathology. Among these kinases, GSK-3β is reported to affect amyloid precursor protein (APP) processing and tau pathology. In 1988, GSK-3β was found to phosphorylate PHF-like epitopes on tau.1 Later studies demonstrated that GSK-3β is a leading candidate kinase responsible for tau hyperphosphorylation in AD. In addition to its roles in tau phosphorylation, GSK-3β is also reported to be involved in APP processing. Aplin et al. showed that GSK-3β is capable of phosphorylating the cytoplasmic domain of APP at Thr743 in vitro. Coexpression of GSK-3β and truncated APP induces an increase in maturation of APP,2 indicating that GSK-3β may regulate APP processing via a phosphorylation mechanism. The relationship between APP processing and GSK-3β is further explained by several recent studies. Morfini et al. reported that GSK-3β colocalizes with membrane-bound organelles and phosphorylates kinesin light chain (KLC) C termini.3 These results indicate that GSK-3β may be directly involved in APP function in neurons because APP is known to bind the C termini of KLC and is subjected to fast anterograde axonal transport.4,5 In 1998, our group first observed that presenilin 1 binds GSK-3β, indicating that GSK3β may regulate amyloid generation of APP processing.6 To study the role of GSK-3β in amyloid generation, we used lithium, a potent GSK-3β inhibitor, to inhibit the activity of GSK-3β, and examined amyloid secretion from cells overexpressing APPC100. Furthermore, we injected fibrillar Aβ into the brains of transgenic mice in the presence or absence of lithium in vivo. We observed that lithium could reduce Aβ secretion in the cells and attenuate the degree of Aβinduced tau pathology in mice.

10.2 EXPERIMENTAL PROCEDURES 10.2.1 LITHIUM TREATMENT

IN

TRANSFECTED CULTURED CELLS

A cDNA (APP C100) encoding 99 amino acids from the C-terminal end of human APP was cloned into the expression vector pCIneo with the cytomegalovirus promoter. For the transient expression of APP C100, COS7 cells were grown and maintained in D-MEM with 10% fetal calf serum (FCS). 1. COS7 cells are seeded into 12 wells with 80 to 90% confluence 1 day before transfection. On the following day, Lipofectamine 2000 (Gibco, now Invitrogen, Carlsbad, CA) is used to perform transfection according to the manufacturer’s instructions. 2. Lithium chloride (LiCl) at a concentration of 1 M is freshly prepared using serum-free medium each time. Final concentrations of 5, 10, and 20 mM LiCl are used in the experiment. 3. Twenty-four hours after transfection, cells are conditioned in serum-free media for 12 or 24 hr in the presence or absence of LiCl. The media are collected for subsequent Aβ measurements.

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10.2.2 Aβ ELISA Sandwich Aβ ELISAs were developed to quantitate Aβx-40 and Aβx-42 as reported previously.7 The end-specific monoclonal antibodies, anti-Aβx-40 (MBC40) and anti-Aβx-42 (MBC42), were used as the capture antibodies. Biotinylated monoclonal anti-N-terminal Aβ secondary antibodies 2H8 and 6E10 were used as the detection antibodies. Streptavidin-conjugated alkaline phosphatase and AttoPhos (Amersham Biosciences, Uppsala, Sweden) were used as the reporter systems. AttoPhos fluorescence was obtained with 444-nm excitation (emission at 555 nm). 1. NUNC Maxisorb immunoassay plates (Nalge NUNC International, Denmark) are coated with 0.3 µg/well (100 µl) capture antibodies in filtered PBS overnight at 4ºC and sealed with sealer. 2. On the next day, plates are blocked with 300 µl/well Block ACE (Dainippon Pharmaceutical, Osaka, Japan, #UK-B80, 1:4 dilution of original solution with deionized water) for 2 hr at room temperature. Wash the plates with PBS-T (PBS with 0.05% Tween-20) once. Load the samples in the wells (100 µl/well). 3. Incubate samples with coated antibodies overnight at 4ºC. Wash with PBS-T twice. 4. The plates are then incubated in a solution of the detector antibody for 2 hr at 4ºC. 5. Wash the plates with PBS-T twice followed by treating the plates with alkaline phosphatase for 1.5 hr (streptavidin-conjugated alkaline phosphatase, Amersham, 1:5000) at 4oC. 6. Wash plates with TBS-T twice. The signal is amplified by adding 100 µl AttoPhos (freshly prepared) and measured with a Fluoroskan (Thermo Labsystems, Vantaa, Finland). Several scans are needed to achieve the best signal. 7. To construct standard curves, Aβ1-40 and Aβ1-42 peptides (AnaSpec, San Jose, CA) are dissolved in dimethyl sulfoxide (DMSO, 1 mg/ml). Further serial dilutions are performed by using Block Ace (1:10 dilution of original solution with 0.05% Tween-20) or 3% BSA with 0.05% Tween-20.

10.2.3 PREPARATION

OF FIBRILLAR

Aβ (FAβ)

1. Aβ1-40 peptide (U.S. Peptide, Rancho Cucamonga, CA) is dissolved in 100% 1,1,1,3,3,3-hexafluoro-2-propanol buffer to a final concentration of 1 mg/ml. 2. Just prior to performing the experiment, aliquots of Aβ are evaporated, redissolved in HEPES buffer (Sigma), and the resulting 5 µM Aβ-40 solution is incubated for 6 hr at 37˚C. 3. Aβ aggregation and toxicity are assessed with Western blotting and ThT, MTT, and histochemical staining as described previously.8

10.2.4 MICROINJECTION

OF FAβ INTO

TAU TRANSGENIC MICE

1. Three-month-old V337M and wild-type (WT) tau transgenic (Tg) mice and non-Tg littermates are anesthetized with 25 mg/kg pentobarbital (Abbott Laboratories, Chicago, IL) and positioned in a stereotaxic apparatus.

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2. An injection needle of a Hamilton syringe is lowered into area CA1 of both hippocampi (coordinates: –1.9 mm from bregma, ±1.0 mm from midline, –1.9 mm dorsoventral). 3. Next, 1 µl of fAβ, sAβ, or polyglycine is injected with a 10-µl Hamilton syringe driven by a mini-pump (Motorized Stereotaxic Injector, Stoelting, Wood Dale, IL) at a rate of 0.05 µl/min. The needle is kept at the injection site for 20 min and then slowly withdrawn. 4. Intrahippocampal circuitry involved in the fAβ injection and the retrograde labeled cells is assessed by stereotaxic injection of 50 nl of fluorogold dextran, a neuronal tracer (Fluoro-Chrome, Englewood, CO) into CA1 as previously described.9 All microinjection operations are carried out on a clean, specialized bench; none of the mice in our tests developed infections.

10.2.5 LITHIUM ADMINISTRATION 1, fAβ-injected V337M Tg mice receive daily intraperitoneal injections (400 µl) of 0.3 M LiCl for 14 or 28 days. Control fAβ-injected V337M Tg mice receive a single 400-µl injection of 0.3 M NaCl. 2. LiCl injections begin 1 day before fAβ injection into area CA1 of the hippocampus, and continue until mice are euthanized. 3. On the final day of treatment, the animals are killed immediately after the beginning of the 12-hr light cycle and analyses are carried out.

10.3 RESULTS 10.3.1 ESTABLISHMENT

OF

SELECTIVE SANDWICH Aβ ELISA

A selective sandwich Aβ ELISA was developed to assay secreted Aβ from cells as described above. To determine the sensitivity and specificity of the assay, the synthetic peptides were diluted with loading buffer in a range from 10 ng/ml to 3.125 pg/ml and loaded into a 96-well plate. Figure 10.1a shows the standard curves obtained from this experiment. Both Aβ-140 and Aβ-142 had a detection limit of 3.125 pg/ml (0.078 fmol/well). No cross-reactivity was observed between Aβ1-40 and Aβ1-42, even with higher concentration loadings of the peptides (10 ng/ml). The specificity of MBC40 and MBC42 was further confirmed in Western blot analysis. Because APP is the precursor of Aβ, the cross-reactivity of these capture antibodies (MBC40 and MBC42) to the full-length and APP C-terminal fragments (CTFs) was investigated. We overexpressed human APP695 and APPC99 in COS7. After immunoprecipitation with an anti-APPC antibody (anti-APPC), the immunoprecipitates of APPexpressing cells were immunoblotted by another anti-APPC antibody (61C), 6E10, MBC40, and MBC42. Both 61C and 6E10 labeled the full-length APP as expected (Figure 10.1b). MBC40 and MBC42 did not label full-length APP. Additionally, the immunoprecipitates of the cells expressing APPC-terminal fragment 99 (APPC100) were immunoblotted with 61C, 6E10, MBC40, and MBC42. The results demonstrated

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a.

b. Aß1-40

30

Aß1-40 Aß1-40 Aß1-42 Aß1-42

R.F.U

25 20 15

2

3

4

1

2

3

4

114

10

84

5 0

0

25

50

75

100

[pg/ml]

17

Aß1-42

Aß1-42 Aß1-42 Aß1-40 Aß1-40

6

R.F.U

1

5 4

8

3 2 1 0 0

25

50

75

100

[pg/ml]

FIGURE 10.1 Sensitivity and specificity of sandwich ELISAs for both Aβx-40 and Aβx-42. (a) Standard curves for Aβ ELISAs. Both anti-Aβ40 and anti-Aβ42 detect the corresponding synthetic peptides at 3.125 pg/ml (0.078 fmol/well). No cross-reactivity was observed even at peptide concentrations up to 10 ng/ml. Replications show duplicates of each peptide loaded in the experiments. (b) The upper panel of the Western blot shows that MBC40 and MBC42 did not label the immunoprecipitated full-length APP from APP-overexpressed COS7 cells. The lower panel of the Western blot shows that MBC40 and MBC42 did not label the immunoprecipitated APP C terminal fragments from APP C100-overexpressed COS7 cells. Lane 1, anti-APP antibody 61C; lane 2, anti-Aβ antibody 6E10; lane 3, MBC40; and lane 4, MBC42.

that 61C detected CTFs of APP and 6E10 detected APPC100 as expected. Again, MBC40 and MBC42 failed to label APPCTFs (Figure 10.1c). The results demonstrated that our Aβ ELISA has a good sensitivity and specificity for selectively detecting Aβ40 and Aβ42.

β SECRETION IN ASSOCIATION WITH GSK-3β β 10.3.2 EFFECT OF LICL ON Aβ ACTIVITY FROM APP C100-TRANSFECTED COS7 CELLS As a first step for determining the regulation of secreted Aβ by GSK-3, APP C100 was transiently expressed in COS7 cells. Western blot analysis of the detergent extract from the transfected cells shows a band with a molecular weight of approximately 14 kDa, corresponding to APP C100 (data not shown). Aβ1-40 and Aβ1-42 were then measured using the selective Aβ sandwich ELISAs. In agreement with the results from the APP C100 expression experiment, abundant secretion of Aβ was detected from the transfected cells. The concentrations of Aβx-40 and Aβx-42, respectively, were 296.39 ± 3.063 pg/ml and 9.071 ± 0.628 pg/ml (data not shown). The percentage of Aβx-42 compared to total Aβ was approximately 5%, demonstrating that most of the Aβ was in the Aβx-40 form.

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a.

b. [pg/ml]

400

Aß1-40

300

*

200

** ***

LiCl 17

100 0

0.

5.

10.

0

5

10

20 (mM)

20.

LiCl (mM)

[pg/ml]

30

Aß1-42

8

20 * 10 0

0.

5.

10.

20.

LiCl (mM)

FIGURE 10.2 Aβx-40 and Aβx-42 secretions in APP C100-transfected cells following LiCl treatment for 24 hr (a) Bar graphs showing dose-dependent reductions of Aβx-40 and Aβx-42. Aβ secretion was measured by sandwich Aβ ELISAs. *p < 0.05, **p < 0.01, ***p < 0.001. (b) Western blot showing slight enhancement of APP C100 expression with increasing LiCl concentration.

Previous studies showed that LiCl inhibits the activity of GSK-3β.10 In order to study the role of GSK-3β in Aβ generation, various concentrations of LiCl were added to APP C100-transfected cells. To demonstrate that any decrease in Aβ secretion was not due to LiCl toxicity resulting in cell death, cell counts were performed at the end of the 24-hr treatment period. No clear cellular toxicity was observed at any LiCl concentration tested (data not shown). LiCl applied at concentrations of 0, 5, 10, and 20 mM decreased both Aβ40 and Aβ42 secretion in a dose-dependent manner (Figure 10.2a). The LiCl-associated decrease in Aβ secretion was statistically significant. The effect of lithium on Aβ secretion was also confirmed in primary neuronal cultures (data not shown). Interestingly, examination of cell lysates revealed a slight, dose-dependent increase in cellular APP C100 as a result of LiCl treatment (Figure 10.2b).

β ON FAβ β-INDUCED TAU PATHOLOGY IN VIVO 10.3.3 EFFECT OF GSK-3β Because we observed that inhibition of GSK-3β reduced Aβ secretion, we further examined inhibition of GSK-3β by lithium on Aβ-induced tau pathology in vivo. In P301L mutant tau Tg mice, fibrillar Aβ42 injection into brains was reported to accelerate the formation of neurofibrillary tangle (NFT)-like tau pathology.11 We injected Aβ into the brains of 3-month-old V337M mutant tau Tg mice (V337M Tg) in which no tau pathology was apparent.12 We found that the injection of fAβ into the CA1 of V337M Tg robustly induced hyperphosphorylated tau in the hippocampi of the mice as demonstrated by biochemical and immunohistochemistry studies (Figure 10.3). Since we previously showed that Aβ-induced tau hyperphosphorylation could be blocked by the inhibition of GSK-3β in hippocampal primary culture,13,14 we studied the effect of inhibition of GSK-3β in tau pathology of Aβ-injected V337M

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a.

b. Vehicle

Aß

Vehicle

Aß

Aß+LiCl

Aß injection

AT180

AT100

Aß injection+LiCl

Aß+LiCl

Tau-C

AT100

SDS-insoluble tau

FIGURE 10.3 (See color insert following page 114.) The effects of GSK-3β inhibition on the Aβ-induced formation of NFT-like tau pathology. (a) Immunohistochemistry study showing that the Aβ-induced tau pathology was blocked in the hippocampus of M337V Tg mice coinjected with Aβ and lithium as compared to mice injected with Aβ alone. (b) Western blot analysis showing that hyperphosphorylated tau was reduced in the hippocampal tissues of mice coinjected with Aβ and lithium as compared to mice injected with Aβ alone.

Tg. Inhibition of GSK-3β by LiCl administration could attenuate the AK-3-induced tau phosphorylation and aggregation (Figure 10.3). Furthermore, inhibition of GSK-3β in these mice can rescue memory loss (data not shown), suggesting that GSK-3β plays an essential role in Aβ-induced pathological and behavioral changes even in vivo.

10.4 DISCUSSION AD patients with different genetic backgrounds exhibit the same neuropathology, namely amyloid senile plaques, NFTs formed by accumulations of paired helical filaments (PHFs), and loss of neurons. Converging evidence suggests that blockades of both amyloid and tau pathologies are key steps in treating this disease. In this chapter, we have described assays for studying amyloid generation and tau phosphorylation. The sandwich Aβ ELISA discussed here shows high sensitivity and specificity. It can serve as an assay to screen the effects of different drugs on Aβ generation and measure endogenous Aβ from human and animal samples. To extend this assay, different combinations of capture antibodies and detector antibodies can be developed to differentiate the roles of several species of Aβ40 and Aβ42 on this disease. The method showing induction of hyperphosphorylated tau by microinjection of fAβ into transgenic mice builds the bridge between amyloid and tau pathology. It is a useful approach to explore the fAβ-induced downstream events in vivo.

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The mechanism underlying the development of both amyloid deposition and tau hyperphosphorylation in AD still is not clear, although we hypothesize that some factors might facilitate both amyloid and tau pathologies. The precise identities of these factors are still unknown. One promising candidate is GSK-3, an important “housekeeping” kinase that plays multiple roles in the central nervous system.15 Particularly, GSK-3β is shown to phosphorylate tau protein in vitro and in vivo and interact with presenilin 1.6,16 We propose then, that under certain pathological conditions, GSK-3β functions as a mediator underlying amyloid overproduction and subsequent tau hyperphosphorylation. For many years, GSK-3β has been well recognized to have the ability to phosphorylate tau protein. The association of GSK-3β with amyloid has not been reported. In 1996, Klein and Melton first reported that lithium potently inhibits GSK-3β activity.10 Additional data from in vitro and intact cell systems show that LiCl specifically inhibits GSK-3β. LiCl at concentrations of 10 to 20 mM reduces tau phosphorylation in cells coexpressing tau and GSK-3β.17 To examine the role of GSK-3β on amyloid secretion, we applied LiCl at concentrations of 5 to 20 mM to APP C100-transfected cells. Decreased Aβ secretion was observed at concentrations as low as 5 mM, a concentration within the optimal range for the inhibition of GSK-3β activity. The results support our hypothesis that GSK-3β is involved in amyloid secretion. Later studies confirmed our findings showing that inhibition of GSK-3β in vivo could eliminate plaque formation in APP Tg mice as well.18 The molecular mechanism underlying the regulation of Aβ secretion by GSK-3β is not clear. The observation that LiCl causes a slight accumulation of APP C100 indicates that GSK-3β might affect γ-secretase activity. This is consistent with the finding that GSK-3β specifically binds presenilin 1 and phosphorylates presenilin.6,19 Alternatively, APP phosphorylated by GSK-3β might have a different accessing ability to the processing enzymes, resulting in altered amyloid generation. In addition to assaying the role of GSK-3β in APP processing, we further extended our study to tau pathology in tau Tg mice. As reported previously in P301L mutant tau Tg mice,11 we observed that Aβ injection accelerated hyperphosphorylated tau and caused neurodegeneration in V337M Tg mice. We found in Aβ-injected V337M Tg mice that activation of GSK-3β was paralleled to tau pathology and associated with accelerated neurodegeneration. Administration of lithium in these mice not only reduced Aβ-induced formation of NFTs, but also rescued synaptic loss, neuronal loss, and subsequent memory impairment. The findings indicate that inhibition of GSK-3β by lithium possibly eliminates amyloid and tau pathologies in AD. In view of the results presented here, several groups report that AD neuropathology is not observed in individuals diagnosed with demented schizophrenia, since lithium is commonly used in the treatment of various psychiatric disorders including schizophrenia.20,21 Chronic administration of lithium with other medications, therefore, may reduce overproduction of amyloid initiated by various factors. Exploring the action of LiCl on GSK-3β should play a significant role in developing clinical therapies for AD.

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REFERENCES 1. Ishiguro, K. et al. A novel tubulin-dependent protein kinase forming a paired helical filament epitope on tau. J. Biochem. (Tokyo) 104, 319–321, 1988. 2. Aplin, A.E. et al. Effect of increased glycogen synthase kinase-3 activity upon the maturation of the amyloid precursor protein in transfected cells. Neuroreport 8, 639–643, 1997. 3. Morfini, G. et al. Glycogen synthase kinase 3 phosphorylates kinesin light chains and negatively regulates kinesin-based motility. EMBO J. 21, 281–293, 2002. 4. Kamal, A. et al. Axonal transport of amyloid precursor protein is mediated by direct binding to the kinesin light chain subunit of kinesin-I. Neuron 28, 449–459, 2000. 5. Koo, E.H. et al. Precursor of amyloid protein in Alzheimer disease undergoes fast anterograde axonal transport. Proc. Natl. Acad. Sci. USA 87, 1561–1565, 1990. 6. Takashima, A. et al. Presenilin 1 associates with glycogen synthase kinase-3beta and its substrate tau. Proc. Natl. Acad. Sci. USA 95, 9637–9641, 1998. 7. Sun, X. et al. Intracellular Abeta is increased by okadaic acid exposure in transfected neuronal and non-neuronal cell lines. Neurobiol. Aging 23, 195–203, 2002. 8. Yoshiike, Y. et al. New insights on how metals disrupt amyloid beta-aggregation and their effects on amyloid-beta cytotoxicity. J. Biol. Chem. 276, 32293–32299, 2001. 9. Zappone, C.A. and Sloviter, R.S. Commissurally projecting inhibitory interneurons of the rat hippocampal dentate gyrus: a colocalization study of neuronal markers and the retrograde tracer fluoro-gold. J. Comp. Neurol. 441, 324–344, 2001. 10. Klein, P.S. and Melton, D.A. A molecular mechanism for the effect of lithium on development. Proc. Natl. Acad. Sci. USA 93, 8455–8459, 1996. 11. Gotz, J. et al. Formation of neurofibrillary tangles in P301l tau transgenic mice induced by Abeta 42 fibrils. Science 293, 1491–1495, 2001. 12. Tanemura, K. et al. Neurodegeneration with tau accumulation in a transgenic mouse expressing V337M human tau. J. Neurosci. 22, 133–141, 2002. 13. Takashima, A. et al. Exposure of rat hippocampal neurons to amyloid beta peptide (25-35) induces the inactivation of phosphatidyl inositol-3 kinase and the activation of tau protein kinase I/glycogen synthase kinase-3 beta. Neurosci. Lett. 203, 33–36, 1996. 14. Takashima, A. et al. Activation of tau protein kinase I/glycogen synthase kinase-3 beta by amyloid beta peptide (25-35) enhances phosphorylation of tau in hippocampal neurons. Neurosci. Res. 31, 317–323, 1998. 15. Grimes, C.A. and Jope, R.S. The multifaceted roles of glycogen synthase kinase 3 beta in cellular signaling. Progr. Neurobiol. 65, 391–426, 2001. 16. Spittaels, K. et al. Glycogen synthase kinase-3beta phosphorylates protein tau and rescues the axonopathy in the central nervous system of human four-repeat tau transgenic mice. J. Biol. Chem. 275, 41340–41349, 2000. 17. Hong, M. et al. Lithium reduces tau phosphorylation by inhibition of glycogen synthase kinase-3. J. Biol. Chem. 272, 25326–25332, 1997. 18. Phiel, C.J. et al. GSK-3 regulates production of Alzheimer’s disease amyloid-β peptides. Nature 423, 435–439, 2003. 19. Kirschenbaum, F. et al. J.V. Glycogen synthase kinase-3 beta regulates presenilin 1 C-terminal fragment levels. J. Biol. Chem. 276, 30701–30707, 2001. 20. Arnold, S.E., Franz, B.R., and Trojanowski, J.Q. Elderly patients with schizophrenia exhibit infrequent neurodegenerative lesions. Neurobiol. Aging 15, 299–303, 1994. 21. Arnold, S.E. et al. Prospective clinicopathologic studies of schizophrenia: accrual and assessment of patients. Am. J. Psychiatr. 152, 731–737, 1995.

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11

Immunocytochemical Analysis of Amyloid Precursor Protein and Its Derivatives Gunnar Gouras and Reisuke H. Takahashi

CONTENTS 11.1 Introduction 11.2 Experimental Approaches 11.2.1 Immunocytochemistry: Fixation 11.2.2 Immunocytochemistry: Controls 11.2.3 Immunocytochemistry for APP and Aβ 11.2.4 Immuno-Electron Microscopy for APP and Aβ 11.3 Conclusion 11.4 Experimental Procedures 11.4.1 Protocol I: Immunostaining Protocol for Paraffin-Embedded Sections 11.4.2 Protocol II: Immunoperoxidase Staining Protocol for Floating Sections 11.4.3 Protocol III: Immunoperoxidase Electron Microscopy 11.4.4 Protocol IV: Immuno-Electron Microscopy with Gold Acknowledgments References

11.1 INTRODUCTION Histochemical analysis of postmortem human brains from subjects who suffered from dementia led to the discovery of Alzheimer’s disease a century ago. The histological lesions that characterized this common age-related dementia were the accumulation of senile plaques (SPs) and neurofibrillary tangles (NFTs). Subsequently, NFTs were found in a variety of degenerative diseases of the brain, while plaques remained more unique to Alzheimer’s dementia. The biochemical analysis of plaques in the 1980s led first to the identification of the fundamental component

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of plaques, the β-amyloid peptide (Aβ),1 followed by the identification of its longer precursor protein, the amyloid precursor protein (APP).2 Since APP was cloned and sequenced, molecular biology studies have provided multiple insights into the biology of APP and Aβ. Histochemical studies have taken a backseat in basic Alzheimer’s research, appearing often as simply descriptive. Few biological studies are wholly accepted before in vivo support is provided by data from the diseased human brain. The combination of antibodies with histology provides a powerful tool to confirm changes in a given protein in a diseased brain. Indeed, immunohistochemistry (or immunocytochemistry, ICC) is similar to a Western blot in that the antibody must find its specific antigen. This is followed by visualization of a linked secondary antibody against the primary antibody via a chemical reaction (chromophore). In contrast to Western blot, where the protein antigen is located on a charged membrane, ICC detects the protein in its native site within the brain. A difficulty in analyzing ICC in neuroscience is that an understanding of the brain cytoarchitecture and anatomy is important. Although viewed by biochemists as less accurate, ICC procedures are more direct in taking the primary antibodies to their respective antigens and avoid some of the intermediate steps required for biochemical procedures such as protein extraction and immunoprecipitation.

11.2 EXPERIMENTAL APPROACHES 11.2.1 IMMUNOCYTOCHEMISTRY: FIXATION ICC generally requires fixation of tissue prior to incubation with the primary antibody. Fixation allows for the preservation of the brain cytoarchitecture, although it can also introduce problems. The fixation procedure can lead to chemical crosslinking which may disrupt the antigenic site of a protein. Trial and error allow for the assessment of optimal ICC conditions for a given antigen. The most common fixative is 4% paraformaldehyde. Weaker fixative procedures such as alcohol-based dehydration with methanol can be employed for antigens that are more sensitive to standard fixation. Stronger fixatives, used especially for electron microscopy (EM) to further preserve the ultrastructural morphology, often include glutaraldehyde with paraformaldehyde. ICC generally requires treatment with a detergent to allow for sufficient penetration of antibodies into the tissue. Triton X-100 is the most common detergent for ICC. No detergent or a lower concentration of Triton or saponin can be used if detergent needs to be minimized (1) because a more lipid-associated antigen would be extracted and thereby lost or (2) in the case of immuno-EM, to avoid damage to the cytoarchitecture that can be observed at an ultrastructural level after treatment with detergent. For ICC and immuno-EM, a balance must be found between the optimal level of fixation and detergent used. As can occur with Western blot, ICC may not adequately detect certain antigens.

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11.2.2 IMMUNOCYTOCHEMISTRY: CONTROLS Potential problems with ICC, as with all antibody-based assays, include potential false cross-reactivity of the antibody and detection of a reaction product not related to the antibody. The latter can be explored by adding excess antigen (such as Aβ peptide) prior to addition of the primary antibody — a technique known as peptide competition. Loss of staining after peptide competition indicates that the antibody sees its antigen, but does not provide information on whether cross-reactivity with an unwanted antigen is present because the antibody is sequestered from the potential cross-reactivity by the excess antigen. The best control for antibody specificity is failure to observe the immunolabeling with a given antibody on knockout mouse tissue that lacks the corresponding antigen.

11.2.3 IMMUNOCYTOCHEMISTRY

FOR

APP

AND

Aβ

With regard to immunostaining of Aβ peptides and APP (full-length and APP C-terminal fragments), an understanding of the epitopes of a given antibody and the processing of APP is important. See Protocols I and II for ICC for Aβ or APP on fixed and frozen and paraffin-embedded brain tissue, respectively. C-terminal-specific APP antibodies can be used to study full-length APP and various C-terminal fragments of APP. N-terminal APP antibodies can be used to study full-length APP and various secreted forms of APP (sAPP). In addition, the significant sequence homology of APP with other members of the APP–APLP (amyloid precursor-like protein) family of proteins (APLP1 and APLP2) may lead to cross-reactivity of some N-terminal APP antibodies with APLPs. The C terminus of APP is less homologous and therefore C-terminal antibodies tend to be specific for APP. One also must be aware when using anti-Aβ antibodies that antibodies raised against the Aβ domain of APP, which includes many of the widely used Aβ antibodies for ICC, also see full-length APP and Aβ-containing APP CTFs (such as βCTF). Moreover, an understanding of the complexity of APP processing allows appreciation of the fact that one may also be assaying various lesser known APP and Aβ fragments. For example, the β-secretase BACE was demonstrated to cleave both at AβAsp1 and at AβGlu11, generating Aβ1-40/42 and Aβ11-40/42 peptides, respectively.3,4 Indeed, the first plaques observed with AD pathology are composed of N-truncated Aβ and these peptides aggregate more readily than full-length Aβ. Therefore when using an antibody against the N terminus of Aβ, one may be missing an important pool of N-truncated Aβ.5 We should note that just because an antibody was raised against a certain Aβ peptide domain, the actual epitope is not necessarily the entire domain. For example, the widely used monoclonal antibody 6E10 (Signet Laboratories, Dedham, MA) against the N terminus of Aβ was raised against Aβ1-16, but has been epitopemapped to Aβ5-10. Thus, 6E10 will not see N-truncated Aβ11-40/42 peptides. A more recent area of research reports that Aβ antibodies recognize more than plaques in AD brains or in transgenic mice that develop plaque pathologies. Specifically,

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* +

* *

FIGURE 11.1 Immunohistochemistry of human AD brain with an Aβ42-specific antibody. Note plaque and intraneuronal Aβ42 staining.

intraneuronal Aβ accumulation by ICC has been observed with AD pathogenesis.6–8 A prerequisite for observing intraneuronal Aβ with concomitant plaque pathology (Figure 11.1) is allowing the immunoperoxidase reaction product enough time to visualize more than the most abundant signal coming from plaques. This is analogous to allowing a Western blot to develop for a sufficient time so that both the full-length APP band and the less abundant Aβ band can be seen when employing an antibody directed against the Aβ domain of APP on cell lysate. Intraneuronal Aβ can be especially appreciated prior to plaque development when the “internal” peptide competition from the high concentration of Aβ in plaques is less, such as in a Down’s syndrome brain destined to develop plaque pathology.6,9,10 An additional complexity is that Aβ antibodies have different affinities, depending on the aggregation state of Aβ. If formic acid is employed to expose even the β-pleated Aβ within plaques, antibodies that prefer monomeric Aβ reveal remarkably more plaque staining. Increasing evidence links Aβ oligomers with the pathogenesis of AD,11 and Aβ oligomer-specific antibodies are being developed to better study Aβ oligomers in brains.

11.2.4 IMMUNO-ELECTRON MICROSCOPY

FOR

APP

AND

Aβ

Generally, milder fixation and detergent conditions are required for immuno-EM than for ICC. After incubation with the primary antibody, staining methods for

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immuno-peroxidase and immuno-gold are different. See Protocols III and IV for details on both methods. With regard to fixation, glutaraldehyde preserves ultrastructural morphology, but at the same time it can prevent the primary antibodies from recognizing antigens because aldehyde residues may block antibody–antigen interactions. To avoid these qualities of glutaraldehyde, some employ acrolein for fixation.12 Acrolein is a milder fixative reagent than glutaraldehyde, especially for neuronal tissues, and does not block antibody–antigen binding as much as glutaraldehyde. Mice or rats are perfused with 3.75% acrolein and 2% paraformaldehyde to preserve the ultrastructure and antigenicity. For human brain, biopsy tissue is rapidly immersion-fixed with 1.875% acrolein and 2% paraformaldehyde. After perfusion, brain tissue is cut (40 µm thick) on a vibrating microtome and treated with sodium borohydride to remove extra aldehyde sites.13,14 A treatment with a detergent is generally required to allow antibodies to penetrate to the antigen in the tissue. Triton X-100 (~0.05%) is commonly used as a detergent for immuno-EM. Freeze–thaw methods can also be used for EM instead of detergent. To allow for penetration of the antibody, tissue is physically destroyed by liquid freon and liquid nitrogen. In some cases, both methods may be required to detect antigen. An optimal combination must be found for the given antibody and antigen. We employed Aβ42 C-terminal-specific antibodies and APP C-terminal-specific antibodies to study localization in brains of Aβ42 and APP, respectively. Employing well-characterized C-terminal Aβ42-specific antibodies MBC42 (kindly provided by Dr. H. Yamaguchi, Gunma University, Gunma, Japan) and AB5078P (Chemicon, Temecula, CA), specific localization of Aβ42 to multivesicular bodies and endosomal vesicles was observed (Figure 11.2).15 In contrast, antibodies to the C terminus of APP predominantly labeled the Golgi apparatus.15,16

FIGURE 11.2 Immuno-gold electron microscopy demonstrating Aβ42 localization especially on the outer limiting membranes of multivesicular bodies in a neuron of a normal mouse.

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11.3 CONCLUSION Immunohistochemical methods are powerful tools for investigating the anatomical distribution of a given antigen, such as APP and its derivatives in brain. Immuno-EM can be especially useful in exploring the cell biology and subcellular neuropathology of Alzheimer’s disease.

11.4 EXPERIMENTAL PROCEDURES 11.4.1 PROTOCOL I: IMMUNOSTAINING PROTOCOL FOR PARAFFINEMBEDDED SECTIONS* Day 1 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17.

Deparaffinize in xylene I, 5 min. Deparaffinize in xylene II, 5 min. Deparaffinize in xylene III, 5 min. 100% ethyl alcohol, 3 min. 100% ethyl alcohol, 3 min. 95% ethyl alcohol, 3 min. 70% ethyl alcohol, 3 min. 50% ethyl alcohol, 3 min. 0.1 M PB, 3 min × 2. 0.1 M Tris-buffered saline (TBS), 3 min × 2. 99% formic acid, 5 min (for optimal plaque detection with Aβ antibodies only). Wash with 0.1 M PBS, 5 min × 3. 3% hydrogen peroxide in 0.1 M TBS, 30 min. Wash with 0.1 M TS, 2 min × 3. Block in 0.5% bovine albumin serum (BSA) in 0.1 M TBS for 30 min (steps 15 through 17 usually performed with 0.01 to 0.03% Triton). Wash once with 0.1 M TS. Incubate with primary antibody in 0.1% BSA at 4°C overnight (in hydration chamber).

Day 2 1. Wash in 0.1 M TS, 10 min ×3. 2. Incubate in secondary antibody (1:400) in 0.1% BSA for 60 min at room temperature. 3. Prepare ABC-HRP (avidin–biotin horseradish peroxidase complex) solution (Vectastain Elite ABC kit): 1 drop of reagent A and 1 drop of reagent B per 2.5 ml of 0.1 M TS; let it sit at least 30 min. 4. Wash in TBS, 10 min × 3. 5. Incubate in ABC-HRP solution for 30 min. * Utilizing Vectastain ABC kit, Vector Laboratories, Burlingame, CA.

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2

6. Wash in 0.1 M TBS, 10 min × 3. 7. Add 5 µl 30% hydrogen peroxide per 50 ml diaminobenzidine (DAB, Sigma-Aldrich, St. Louis, MO) solution (22mg/100ml). 8. Develop in and time the DAB reaction; check for staining under a light microscope (or use kit from Vector). 9. Rinse 1 min with distilled water. 10. Wash once with 0.1 M TBS. 11. Wash with 0.1 M PB. 12. Place in dessicator, 30 min. 13. Dehydrate in series of increasing alcohol concentrations (50, 75, and 95%) for 3 min each; 100% for 5 min × 2, Xylene I, Xylene II, and Xylene III for 10 min each. 14. Coverslip with DPX mounting medium and let dry overnight.

11.4.2 PROTOCOL II: IMMUNOPEROXIDASE STAINING PROTOCOL FOR FLOATING SECTIONS* Day 1 1. Wash with 0.1 M PBS, 5 min × 3. 2. 99% formic acid, 5 min (for optimal plaque detection with Aβ antibodies only). 3. Wash with 0.1 M PBS, 5 min × 3 4. 3% hydrogen peroxide in 0.1 M PBS, 30 min. 5. Wash with 0.1 M PBS, 5 min × 3. 6. Block in 0.5% BSA in 0.1 M PBS for 30 min (steps 6 through 8 usually performed with 0.01 to 0.03% Triton). 7. Wash once with 0.1 M TBS. 8. Incubate with primary antibody in 0.1% BSA overnight at 4ºC. Day 2 1. Wash with 0.1 M PBS, 5 min × 3. 2. Incubate in secondary antibody (1:400) in 0.1% BSA for 60 min at room temperature. 3. Prepare ABC-HRP (horseradish peroxidase) solution (Vectastain Elite ABC kit) (1 drop of reagent A and 1 drop of reagent B per 2.5 ml of 0.1 M PBS); let sit at least 30 min. 4. Wash in 0.1 M PBS, 5 min × 3. 5. Incubate in ABC-HRP solution for 30 min. 6. Wash in 0.1 M PBS, 5 min × 3. 7. DAB solution: 5 ml dH2O, 2 drops solution, 4 drops DAB solution, 2 drops hydrogen peroxide mixed well (or make DAB solution as in Protocol I). 8. Time DAB reaction and check for staining under light microscope. 9. Wash once with 0.1 M PBS. * Utilizing Vectastain ABC kit, Vector Laboratories, Burlingame, CA.

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10. Mount sections on coated glass slices; let sections dry. 11. Dehydrate in series of increasing alcohol concentrations (50, 75, and 95%) for 3 min each; 100% for 5 min × 2, Xylene I, Xylene II, and Xylene III for 10 min each. 12. Coverslip with DPX.

11.4.3 PROTOCOL III: IMMUNOPEROXIDASE ELECTRON MICROSCOPY Day 1 1. Perfusion fixation with 3.75% acrolein/2% paraformaldehyde in 0.1 M PB; for brain biopsy, fix tissue with 1.875% acrolein/2% paraformaldehyde in 0.1 M PB. 2. Cut brain tissue with Vibratome (40 µm). 3. Wash in 0.1 M PB. 4. 1% sodium borohydride in 0.1 M PB for 30 min. 5. Wash in 0.1 M PB until bubbling stops. 6. Wash in 0.1 M TBS, 10 min × 2. 7. Block in 0.5% BSA/0.1 M TS for 30 min. 8. Wash in 0.1 M TBS, 1 min × 3. 9. Incubate sections with primary antibody in 0.1% BSA/0.1 TS overnight at 4ºC and for a second night at room temperature. Add Triton X-100 (~0.05%) if needed. Day 2 1. Wash in 0.1 M TBS, 10 min × 3. 2. Incubate with secondary antibody (1:400) in 0.1% BSA/0.1 M TBS for 30 min. 3. Prepare ABC-HRP (horseradish peroxidase) solution (Vectastain Elite ABC kit); 2 drops of reagent A and 2 drops of reagent B per 10 ml of 0.1 M TBS; shake immediately and let it sit at least 30 min. 4. Wash in 0.1 M TBS, 10 min × 3. 5. Incubate in ABC solution for 30 min. 6. Wash in 0.1 M TBS, 10 min × 3. 7. During washes, prepare DAB solution and add 22 mg DAB to 100 ml 0.1 M TBS and stir. Just before using, add 10 µl of 30% hydrogen peroxide. 8. Time DAB reaction and check for staining under light microscope. 9. Wash in 0.1 M TBS, 2 min × 2. 10. Wash in 0.1 M PB, 2 min × 3. 11. Carefully lay sections flat in shallow wells containing 0.1 M PB and replace with 2% osmium tetroxide in 0.1 M PB. Incubate 1 hr. 12. Wash in 0.1 M PB, 3 min × 3. 13. 30% ETOH, 5 min. 14. 50% ETOH, 5 min.

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15. 16. 17. 18. 19.

70% ETOH, 5 min. 95% ETOH, 5 min. 100% ETOH, 10 min × 2. Propylene oxide 10 min × 2. Propylene oxide:EPON (mixed 1:1) rotate overnight at room temperature.

Day 3 1. Fresh EPON; rotate 2 hr. 2. Flat embedding at 60ºC for 18 to 24 hr. Embed sections between two sheets of aclar fluorhalocarbon film. Transfer as little EPON as possible to the sheet with the section, and vigorously squeeze the sections together, removing air bubbles and pushing EPON away.

11.4.4 PROTOCOL IV: IMMUNO-ELECTRON MICROSCOPY

WITH

GOLD

Day 1 1. Same as Protocol III. 2. Concentration of primary antibody is generally three to four times higher than for immunoperoxidase method. Day 2 1. Wash once in 0.01 M PBS. 2. Block in washing buffer (0.1% gelatin, 0.8% BSA in 0.01 M PBS, pH 7.4) 10 min. 3. Incubate with gold-conjugated IgG in washing buffer (1:50) for 2 hr. 4. Wash in washing buffer, 5 min. 5. Wash in 0.01 M PBS 5 min × 3. 6. Incubate in 2% glutaraldehyde in 0.01 M PBS for 10 min. 7. Wash in PBS. 8. Wash in 0.2 M citrate buffer (0.2 M citrate sodium, pH 7.4). During washing, mix silver reagents A and B (Amersham Biosciences, Uppsala, Sweden) at a 1:1 ratio. 9. Silver intensification reaction; check for staining under a light microscope. 10. Wash in citrate buffer to stop reaction. 11. Carefully lay sections flat in shallow wells containing 0.1 M PB. 12. Incubate with 2% osmium tetroxide in 0.1 M PB for 1 hr. 13. Wash in 0.1 M PB, 3 min × 3. 14. 30% ETOH, 5 min. 15. 50% ETOH, 5 min. 16. 70% ETOH, 5 min. 17. 95% ETOH, 5 min. 18. 100% ETOH, 10 min × 2. 19. Propylene oxide, 10 min × 2. 20. Propylene oxide:EPON (mixed 1:1); rotate overnight at room temperature.

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Day 3 1. Same as Protocol III.

ACKNOWLEDGMENTS We thank Dr. Noel Calingasan, Department of Neurology and Neuroscience, Weill Medical College of Cornell University, for reviewing this chapter.

REFERENCES 1. Glenner, G.G. and Wong, C.W. Alzheimer’s disease and Down’s syndrome: sharing of a unique cerebrovascular amyloid fibril protein. Biochem. Biophys. Res. Commun. 122, 1131, 1984. 2. Kang, J. et al. The precursor of Alzheimer’s disease amyloid A4 protein resembles a cell-surface receptor. Nature 325, 733, 1987. 3. Vassar, R. et al. Beta-secretase cleavage of Alzheimer’s amyloid precursor protein by the transmembrane aspartic protease BACE. Science 286, 735, 1999. 4. Cai, H. et al. BACE1 is the major beta-secretase for generation of Abeta peptides by neurons. Nat. Neurosci. 4, 233, 2001. 5. Gouras, G.K. et al. Generation and regulation of beta-amyloid peptide variants by neurons. J. Neurochem. 71, 1920, 1998. 6. Gouras, G.K. et al. Intraneuronal Abeta42 accumulation in human brain. Am. J. Pathol. 156, 15, 2000. 7. D’Andrea, M.R. et al. Consistent immunohistochemical detection of intracellular beta-amyloid 42 in pyramidal neurons of Alzheimer’s disease entorhinal cortex. Neurosci. Lett. 333, 163, 2002. 8. D’Andrea, M.R. et al. The use of formic acid to embellish amyloid plaque detection in Alzheimer’s disease tissues misguides key observations. Neurosci. Lett. 342, 114, 2003. 9. Gyure, K.A. et al. Intraneuronal abeta-amyloid precedes development of amyloid plaques in Down syndrome. Arch. Pathol. Lab. Med. 125, 489, 2001. 10. Busciglio, J. et al. Altered metabolism of the amyloid beta precursor protein is associated with mitochondrial dysfunction in Down’s syndrome. Neuron 33, 677, 2002. 11. Selkoe, D.J. Nature, 426, 900, 2003. 12. Leranth, C. and Pickel, V.M. Folding proteins in fatal ways, in Neuroanatomical Tract Tracing Methods, Vol. 2, Plenum, New York, 1989, p. 120. 13. Milner T.A. et al. Hippocampal 2a-adrenergic receptors are located predominantly presynaptically but are also found postsynaptically and in selective astrocytes. J. Comp. Neurol. 395, 310, 1998. 14. Chan, J., Aoki, C., and Pickel, V.M. Optimization of differential immunogold–silver and peroxidase labeling with maintenance of ultrastructure in brain sections before plastic embedding. J. Neurosci. Methods 33, 113, 1990. 15. Takahashi, R.H. et al. Intraneuronal Alzheimer abeta42 accumulates in multivesicular bodies and is associated with synaptic pathology. Am. J. Pathol. 161, 1869, 2002. 16. Caporaso, G.L. et al. Morphologic and biochemical analysis of the intracellular trafficking of the Alzheimer beta/A4 amyloid precursor protein. J. Neurosci. 14, 3122, 1994.

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12

Pathological Detection of Aβ and APP in Brain Chica Mori and Cynthia A. Lemere

CONTENTS 12.1 Introduction 12.2 Methods 12.2.1 Tissue Processing Protocols 12.2.1.1 Paraffin Tissue 12.2.1.2 Frozen Tissue 12.2.2 Immunohistochemistry 12.2.2.1 Paraffin Sections 12.2.2.2 Frozen Sections 12.2.3 Congo Red 12.2.4 Thioflavin S Staining Protocol 12.2.5 Double Immunofluorescent Labeling 12.2.6 Double Labeling with DAB and Other Colored Markers 12.3 Results and Discussion 12.3.1 Aβ40 vs. Aβ42 in Down’s Syndrome 12.3.2 Aβ40 vs. Aβ42 in Familial Alzheimer’s Disease (FAD) 12.3.3 Intraneuronal Aβ in Down’s Syndrome 12.3.4 Intraneuronal Aβ in APP Transgenic Mice 12.3.5 APP Immunoreactivity 12.3.6 Colocalization of Aβ with Glia References.

12.1 INTRODUCTION In 1906, a German physician named Alois Alzheimer described the two major pathologic features, amyloid plaques and neurofibrillary tangles (NFTs), found in the brain of an elderly woman with a dementia-causing neurodegenerative disorder that later became known as Alzheimer’s disease (AD). Subsequently, various histological dyes and stains including silver were used to identify these lesions in autopsied brain tissue. In 1984, Glenner and Wong purified and chemically identified amyloid-β protein (Aβ), a 4-kD monomer, from meningeal blood vessels.1 Later, the sequence was extended from 24 to 40 amino acids2 and found to be identical to the amino acid compositions of senile plaques purified from human AD brains.3,4 0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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FIGURE 12.1 (See color insert following page 114.) Detection of fibrillar amyloid by thioflavin S and Congo red staining. (a) Thioflavin S dye binds to β-pleated Aβ in plaques (arrowheads) and blood vessels (arrows), as demonstrated in this formalin-fixed paraffin brain section of an 18-month-old APP transgenic mouse. Thioflavin S staining is visualized using a fluorescent microscope. Magnification ×10. (b) Congo red also binds fibrillar amyloid and produces a birefringence that alternates between yellow and green under polarized light. Plaques (arrowheads) and vascular Aβ (arrows) are observed in a paraffin section of temporal cortex from a human AD brain. Magnification ×20.

In the 1990s, several mutations in the amyloid precursor protein (APP) gene were correlated with early onset AD5,6 or hereditary cerebral hemorrhage7,8 in a small number of families, setting the stage for the prominent roles of APP and Aβ in AD pathogenesis. In addition, persons with Down syndrome (DS), trisomy 21, develop AD-like pathology, including plaques and NFTs, due primarily to the gene dosage effect of having three copies of the APP gene encoded on chromosome 21.9 Prior to the mid-1980s, plaque and vascular amyloid were identified in brain sections using silver stains or dyes that bound β-sheet amyloid fibrils, such as Congo red and thioflavin S (Figure 12.1). Upon the identification of the Aβ amino acid sequence, Aβ synthetic peptides were manufactured and Aβ-specific antibodies produced by injecting synthetic Aβ peptide into rabbits to generate polyclonal antibodies (Pab) or into mice to generate monoclonal antibodies (Mab). Consequently, Aβspecific antibodies proved exceptionally sensitive for immunohistochemically detecting Aβ protein on brain tissue sections. As a result, the extent of Aβ deposition in aged humans and in particular, AD patients, turned out to be much greater than that previously observed using histological methods. One reason for this unveiling of Aβ deposition was that the some Aβ antibodies were able to identify more than only fibrillar amyloid; they also recognized nonfibrillar (i.e., diffuse, prefibrillar) Aβ deposits. The objective of this chapter is to provide extensive technical details for the preparation of brain tissue sections and the identification of Aβ protein and its precursor, APP, by various histological and immunohistochemical (IHC) methods. These methods may be applied to autopsied human, mouse or monkey brains although the cross-reactivities of certain antibodies should be checked for each species. In addition to specific protocols for immunohistochemistry and histological stains, examples of how they were used in our lab are illustrated. Table 12.1 lists the antibodies we used in immunohistochemistry studies.

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TABLE 12.1 Immunohistochemistry Antibodies Used in Authors’ Laboratory Antibody R1282 (Aβ) 6E10 (Aβ) 21F12 (Aβ42) C42 (Aβ42) BC42 (Aβ42) MBC42 (Aβ42) QCB42 (Aβ42) 2G3 (Aβ40) BC40 (Aβ40) MBC40 (Aβ40) QCB40 (Aβ40) N3pE (Aβ pyroglut 3) N1D (Aβ1-5) 3D6B (Aβ1-5) 8E5 (APP 444-592) C8 (APP, C terminus) 22C11 (APP, N terminus) AT8 (PHF tau) GFAP (human astrocyte) GFAP (mouse astrocyte) CD45 (microglia) HLA-DR (microglia)

Mab or Pab

Source

Dilution

Pretreatment

Pab Mab Mab Pab Pab Mab Pab Mab Pab Mab Pab Pab Pab Mab Mab Pab Mab Mab Pab Mab Mab Mab

Selkoe (Boston, MA) Signet (Dedham, MA) Elan Corp. (Dublin, Ireland) T.C. Saido (Tokyo, Japan) H. Yamaguchi (Gunma, Japan) H. Yamaguchi BioSource (Camarillo, CA) Elan Corp. H. Yamaguchi H. Yamaguchi BioSource T.C. Saido T.C. Saido Elan Corp. Elan Corp. Selkoe Chemicon (Temecula, CA) Innogenetics (Ghent, Belgium) Dako (Carpenteria, CA) Sigma (St. Louis, MO) Serotec (Raleigh, NC) NeoMarkers (Fremont, CA)

1:1000 1:1000 1:1000 1:250 1:500 1:1000 1:100 1:1000 1:500 1:1000 1:100 1:200 1:150 1:200–500 1:1000 1:1000 1:200 1:50 1:1000 1:500 1:5000 1:100

FA FA FA FA FA FA FA FA FA FA FA FA, FA/MW FA, FA/MW FA, FA/MW MW MW MW MW — — MW MW

Mab = monoclonal antibody. Pab = polyclonal antibody. FA = formic acid. MW = microwave.

12.2 METHODS 12.2.1 TISSUE PROCESSING PROTOCOLS Fresh tissue must be fixed, dehydrated, and infiltrated with paraffin prior to paraffin sectioning. Tissue for cryosectioning should be embedded immediately in ornithine carbamyl-transferase (OCT, Sakura Finetek, Torrance, CA) or fixed and then embedded in OCT and frozen at –80ºC. This section provides information about tissue processing and slide preparation for immunohistochemistry. 12.2.1.1 Paraffin Tissue 12.2.1.1.1 Humans Fresh postmortem human brain is cut into small blocks of tissue and then fixed in 10% neutral buffered formalin (pH 7.0 to 7.6) for 1 hr or more at room temperature. Tissue specimens are thoroughly washed in Tris buffer solution (TBS; 50 mM Tris, 150 mM NaCl) to remove excess formalin, placed in cassettes, and stored in TBS at 4ºC until processing.

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The specimens are dehydrated through graded ethanol, cleared of lipids in Histosolve (Shandon, Pittsburgh, PA), and infiltrated with two changes of hot paraffin (~60ºC; temperature varies among brands). The paraffin-saturated specimens are placed into paraffin-containing embedding cassettes, cooled, and stored at room temperature for sectioning. Paraffinized specimens are sectioned 10 to 12 microns thick and placed on glass slides. Slides are dried overnight, baked at 60ºC for 1 hr, and stored at room temperature for immunohistochemistry studies. 12.2.1.1.2 Mice Perfuse mice in 4% paraformaldehyde (PFA), remove brain, and place it in PFA. Follow step 2 and 3 in human tissue protocol. 12.2.1.2 Frozen Tissue Fresh brain specimens (mouse and human) are embedded in OCT. Mouse tissues can be fixed in sucrose or formaldehyde before OCT embedding. They are snapfrozen in liquid nitrogen, then stored at –80ºC for cryosectioning. Specimens should be stored at –20ºC prior to sectioning. They should be sectioned 10 microns thick at –20ºC and placed on glass slides. The slides should be stored at –20ºC for immunohistochemistry studies. Long-term storage should be at –80ºC.

12.2.2 IMMUNOHISTOCHEMISTRY Immunohistochemistry (IHC) studies are used to visualize proteins. Generally, paraffin sections produce better protein morphology with immunohistochemistry than they do in frozen sections. However, formalin fixation prior to paraffin embedding of tissues can mask the antigen and may require antigen retrieval via boiling or with enzyme or acid digestion. These processes may expose antigen, but they can also make the sections brittle. In general, pretreating paraffin sections with formic acid prior to staining (see below) enhances Aβ immunoreactivity. Microwave pretreatment enhances tau, neurofilament and APP immunoreactivity. Frozen sections do not require such a step and surface antigens are well preserved compared to formalin fixed sections. Hence, cytokine staining works better in frozen sections than in paraffin sections. The disadvantages of frozen sections are poor antigen morphology and the need for a freezer that can provide storage at –80ºC. This section provides general protocols for immunohistochemistry studies. Protocols for antigen-enhancing pretreatments are also provided. 12.2.2.1 Paraffin Sections 1. Deparaffinize first in two changes (3 min each) of Histoclear (National Diagnostics, Atlanta, GA) followed by 3 min in a 50:50 mixture of Histoclear and 100% ethanol, and then rehydrate in a graded series of ethanol concentrations (95, 75, and 50% ethanol) ending in water.

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2. Quench endogenous peroxidase activity in 0.3% hydrogen peroxide in methanol for 8 min (99 ml methanol and 1 ml 33% H2O2). Pour out mixture. Rinse twice in distilled water for 3 min each time. 3. Perform antigen-enhancing pretreatment if necessary. Microwave pretreatment — Dilute Antigen Retrieval Citra (BioGenex, San Ramon, CA) ×10 with super-Q water. Put slides into a glass dish filled with Citra. Place the dish in a plastic box filled with 1 in water. Microwave and bring to a boil, then perform cyclic boiling for an additional 5 min. Cool to room temperature. Formic acid pretreatment — Add 88% formic acid (Fisher, Fair Lawn, NJ) onto sections for 10 min. Wash twice in distilled water for 3 min each time. 4. Block in 10% serum (from the animal species in which the secondary antibody was generated) for 20 min at room temperature or 4°C overnight. Dilute serum in TBS. 5. Incubate sections in primary antibodies for 1 hr at room temperature or 4ºC overnight. 6. Wash gently in TBS. 7. Incubate sections in secondary antibodies (goat anti-rabbit for polyclonal antibodies and horse anti-mouse for monoclonal antibodies; Vector Laboratories, Burlingame, CA) for 30 min at room temperature (1 ml serum [goat for polyclonal, horse for monoclonal], 9 ml TBS, and 45 µl secondary antibody). 8. Gently wash in TBS. 9. Incubate in horseradish peroxidase–avidin (Elite ABC kit, Vector Laboratories) 30 min at room temperature (10 ml TBS, 90 µl solution A and 90 µl solution B). 10. Gently wash in 50 mM Tris. 11. Develop in diaminobenzidine (DAB; Sigma Immunochemicals, St. Louis, MO). Dissolve 2 mg DAB into 100 ml of 50 mM Tris; add 33 µl of 33% H2O2 to activate. 12. Gently wash in distilled water. 13. Counterstain with hematoxylin; differentiate with acid alcohol (1 ml concentrated HCl per 100 ml 70% ethanol). 14. Dehydrate in distilled water, 50% ethanol, 75% ethanol, 95% ethanol, two changes of 100% ethanol, 50:50 mixture of 100% ethanol and Histoclear, and two changes of Histoclear. 15. Coverslip with Permount (Fisher Scientific, Pittsburgh, PA). 12.2.2.2 Frozen Sections 1. Air dry the slides for 15 min at room temperature. 2. Add a drop of acetone (stored at –20ºC) or methanol to a section and let it air dry for 15 min at room temperature. 3. Place in TBS for 5 min. 4. Follow paraffin sectioning protocol (12.2.2.1) and continue.

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12.2.3 CONGO RED Congo red recognizes the beta sheet conformations of Aβ fibrils. Images of Congored stained sections are observed using polarized light microscopy. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Deparaffinize and rehydrate. Stain in Mayer’s hematoxylin for 10 min. Wash quickly in distilled water. Differentiate with acid alcohol. Wash in three changes of super-Q water. Wash in freshly alkalinized sodium chloride solution (100 ml saturated NaCl in 80% ethanol and 1 ml 1% NaOH) for 20 min. Filter Congo red (100 ml saturated NaCl in 80% ethanol, 0.2 g Congo red, and 1 ml aqueous NaOH) and use it to stain within 15 min. Place in three changes of 100% ethanol. Dehydrate slides in graded concentration of ethanol and Histoclear. Coverslip with Permount.

12.2.4 THIOFLAVIN S STAINING PROTOCOL Thioflavin S (Sigma, St. Louis, MO) is a dye that binds beta-pleated fibrils. It detects fibrillar amyloids in plaques and blood vessels. Sections are observed under a fluorescent microscope. 1. Deparaffinize tissue in Histoclear and rehydrate in graded ethanol to water. 2. Rinse three times in distilled water. 3. Incubate in filtered (filter fresh before each use) 1% aqueous thioflavin S for 8 min at room temperature (2 g thioflavin S in 200 ml distilled water). 4. Decant thioflavin S and wash for 3 min in 80% ethanol. 5. Decant ethanol and wash again for 3 min in 80% ethanol. 6. Decant ethanol and wash for 3 min in 95% ethanol. 7. Wash with three changes of distilled water. 8. Coverslip in aqueous mounting media (Hydromount, National Diagnostics, Atlanta, GA).

12.2.5 DOUBLE IMMUNOFLUORESCENT LABELING Double immunofluorescent labeling is used to visualize two different proteins in a single section. Signals are detected using a fluorescent microscope. For example, one antigen may be detected as red and the other as green. When two antigens colocalize, the signal appears yellow. This technique is helpful in determining the spatial relationship of two proteins. It also allows us to visualize cellular ingestion of proteins such as when microglia phagocytose Aβ. Day 1 1. Deparaffinize and rehydrate. 2. Apply any pretreatments.

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3. Immerse sections in 0.1 M Tris buffer (pH 7.6) for 5 min (20 ml 1 M Tris and 180 ml distilled water). 4. Block in 2% goat serum (serum from the animal species in which the secondary antibody was generated) in 0.1 M Tris for 5 min (200 µl serum in 9.8 ml Tris buffer). 5. Prepare antibodies. For double labeling, combine the two primary antibodies 50:50 in volume. Dilute antibodies in 2% serum (same species as used in step 4) in 0.1 M Tris. 6. Flick off and wipe off excess fluid and apply antibodies. 7. Incubate overnight (16 to 20 hr) at 4°C in humidified chamber. Day 2 1. Warm slides to room temperature. 2. Rinse sections with 0.1 M Tris. 3. Immerse sections in 2% goat serum (serum from the animal species in which the secondary antibody was generated) in 0.1 M Tris for 5 min. 4. Dilute secondary antibodies (Molecular Probes, Laiden, OR). [Combination 1 = AlexaFluor 488 goat anti-rabbit (green) and rhodamine red goat anti-mouse (red). Combination 2 = AlexaFluor 488 goat anti-mouse (green) and rhodamine red goat anti-rabbit (red)] 5. Centrifuge the mixture of red and green secondary antibodies for 30 sec. Apply the supernatant (1.5 µl AlexaFluor 488, 1.5 µl rhodamine red in 997 µl of 2% serum in 0.1 M Tris). 6. Incubate at room temperature for 2 hr. 7. Rinse secondary antibodies from sections with 0.1 M Tris. 8. Apply Sudan black B for 10 min at room temperature. Make fresh batch if Sudan black is more than 2 months old. Add 0.3 g (0.3%) Sudan black B to 100 ml 70% ethanol. Wrap foil around container, add liquid, powder and a stirring bar. Stir container wrapped in foil for 2 hr using automixer. Store at 4ºC. 9. Rinse sections three times with 0.1 M Tris. 10. Wash in distilled water. 11. Fix in 10% formalin for 1 hr at room temperature. Wrap foil around glass container and its lid. 13. Wash twice in distilled water, 10 min each time. 14. Coverslip with Hydromount. Apply clear nail polish around the edge of the coverslip to prevent sections from drying out. Put slides in closed folder at 4°C.

12.2.6 DOUBLE LABELING

WITH

DAB

AND

OTHER COLORED MARKERS

The principle for DAB labeling is the the same as for immunofluorescent double labeling in that each antigen is detected as a different color and this allows assessment of colocalization of two different antigens. Typically, one antigen is stained by DAB (brown) and the other with an alkaline phosphatase (AP) substrate kit (Vector). The kit allows a choice of red, blue or black. The order of staining is irrelevant in many

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cases although one particular order of staining may be better than another for certain antigens. In general, DAB staining can be followed by AP staining and vice versa. There are four possible combinations: 1. 2. 3. 4.

DAB (antigen 1) followed by AP (antigen 2) DAB (antigen 2) followed by AP (antigen 1) AP (antigen 1) followed by DAB (antigen 2) AP (antigen 2) followed by DAB (antigen 1)

It is important to determine which procedure works best for the antigens of interest. In the procedure below, AP staining is followed by DAB staining. Day 1 1. Deparaffinize and rehydrate. 2. Quench endogenous peroxidase activity in 0.3% hydrogen peroxide in methanol for 8 min. Pour out mixture. Rinse twice in distilled water, 3 min each time. 3. Perform antigen-enhancing pretreatments. 4. Block with 10% serum (from the animal species in which the secondary antibody was generated) in TBS. 5. Apply primary antibody and incubate 1 hr at room temperature. 6. Gently wash in TBS. 7. Apply 45 µl secondary antibody to 10 ml of 10% serum in TBS; incubate 30 min at room temperature. 8. Gently wash in TBS. 9. Prepare ABC for AP staining (45 µl solution A and 45 µl solution B) in 5 ml TBS. Allow to sit 20 min. 10. Apply ABC onto sections and incubate 30 min at room temperature. 11. Gently wash in Tris. 12. Prepare AP substrate kit in your color of interest (red, blue, or black). A. 5 ml of 100 mM Tris at pH 8.2. It is very important to adjust the pH between 8.2 and 8.5 to achieve development. B. Add a drop of levamisole. Shake well. C. Add 90 µl of solution 1. Shake well. D. Add 90 µl of solution 2. Shake well. E. Add 90 µl of solution 3. Shake well. 13. Rinse under cold running water to stop the reaction from proceeding further. 14. Perform antigen-enhancing pretreatments. 15. Block with 10% serum in TBS. 16. Apply the other primary antibody. Incubate overnight at 4°C. If you are in a hurry, you can incubate 1 hr at room temperature and continue on. Day 2 1. Warm slides to room temperature if they have been incubated overnight at 4°C. 2. Gently wash in TBS.

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3. Apply secondary antibody (45 µl secondary antibody in 10 ml of 10% serum in TBS); incubate 30 min at room temperature. 4. Prepare ABC for DAB development (45 µl solution A and 45 µl solution B in 5 ml TBS). Allow to sit 20 min. 5. Apply ABC and incubate 30 min at room temperature. 6. Gently wash in Tris. 7. DAB development. 8. Counterstain with hematoxylin and differentiate with acid alcohol. 9. Dehydrate. 10. Coverslip with Permount.

12.3 RESULTS AND DISCUSSION β40 12.3.1 Aβ

VS.

β42 Aβ

IN

DOWN’S SYNDROME

Down syndrome (DS), trisomy 21, results in an extra copy of the APP gene located on chromosome 21. Most DS patients develop full-blown AD pathologies by the age of 40 or 50.10,11 Plaque deposition begins decades earlier than it does in nonDS individuals. Thus, DS provides a model for studying the temporal progression of the neuropathogenesis of AD. The generation of Aβ C-terminal-specific antibodies (recognizing Aβ42 and Aβ40, specifically) has allowed further insight into the sequence and morphological characterization of the deposition of different species of Aβ. Multiple reports have demonstrated that Aβ42 is deposited into plaques prior to Aβ40. For example, extracellular accumulation of Aβ was reported in diffuse Aβ42 immunoreactive plaques as soon as 12 years of age in DS brain; Aβ40 immunolabeling was not observed (Figure 12.2a and Figure 12.2b).12,13 At older ages, DS brain showed increased numbers of plaques (Figure 12.2c and Figure 12.2d), aggregation of Aβ and compaction of plaques that contained Aβ40 in addition to Aβ42 (Figure 12.2e, Figure 12.2f, Figure 12.3a, and Figure 12.3b).12,14 Biochemical studies of vascular amyloid reported the Aβ deposits contained predominantly Aβ ending at valine 40.15 IHC studies showed that blood vessels in young DS patients had weak Aβ42 immunoreactivity in smooth muscle cells but with increasing age, vascular Aβ deposits were composed mainly of aggregated Aβ40 with some colocalization of lesser amounts of Aβ42 (Figure 12.4).

β40 12.3.2 Aβ

VS.

β42 Aβ

IN

FAMILIAL ALZHEIMER’S DISEASE (FAD)

Although 80 to 90% of total Aβ consists of Aβ40 and only 10 to 20% is Aβ42, Aβ42 is selectively deposited first in senile plaques and is more prone to fibrillization than Aβ40. In addition, Aβ42 aggregates into insoluble amyloid fibrils much faster than Aβ40.16 A study of Colombian patients with early onset FAD caused by an E280A mutation in the presenilin 1 (PS1) gene showed that abundant Aβ42 deposition and gliosis occurred about 30 years earlier than in sporadic AD patients.17 In the PS1-FAD patients, robust increases of Aβ42 were observed in plaques and blood vessels in cortex, hippocampus, and cerebellum, but Aβ40 was not increased relative to that found in sporadic AD cases (Figure 12.5).

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FIGURE 12.2 Aβ C-terminal immunostaining in young DS brain. Antibody BC42 that specifically recognizes residue 42 at the C terminus of Aβ revealed many diffuse plaques (arrowheads) in brains of 12-year-old (a), 17-year-old (c), and 29-year-old (e) DS subjects. Plaques were not detected in adjacent sections with antibody BC40 that specifically recognizes residue 40 at the Aβ C terminus at ages 12 (b) and 17 (d), whereas a small number of plaques were detected at age 29 (f). In addition to the many discrete plaques observed in the 12- and 17-year-old brains, larger patches of Aβ deposition (arrowhead in e) were detected at age 29 by BC42. BC40 darkly stained the cores of a small number of plaques (large arrowheads in f) in the 29-year-old brain. Weak BC40 reactivity (small arrowheads in f) was also detected by BC40 in occasional plaques that stained intensely with BC42 (e). Bar = 200 µm. (Reprinted from Lemere C.A. et al., in Neurobiology of Disease, Vol 3, 1996, p. 19. With permission from Elsevier.)

Over 90 different PS-1 mutations have been cited in the literature. The recently identified early-onset PS-1 mutations, Y256S and Q222H, also exhibited elevated Aβ1-42 levels in brain homogenates. In particular, the PS1 mutation Y256S, with

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FIGURE 12.3 Colocalization of BC42 and BC40 immunostaining within a plaque. Adjacent 8-µm sections of temporal cortex from a 47-year-old DS patient were immunostained with Aβ C-terminal antibodies BC42 (a) and BC40 (b). Bar = 25 µm. (Reprinted from Lemere, C.A. et al., in Neurobiology of Disease, Vol 3, 1996, p. 26. With permission from Elsevier.)

an age of onset at 25 years, showed markedly high Aβ42 levels and some increase in Aβ40 levels.18 The other PS1 mutation, Q222H, with an average age of onset at 35 years, showed an elevation in Aβ42 but not Aβ40. In numerous studies in both human FAD and in APP and PS1/APP transgenic mice, the level of Aβ42 in the brain seemed to correlate with severity of AD in neuropathology and age of onset.

β 12.3.3 INTRANEURONAL Aβ

IN

DOWN’S SYNDROME

Although the origin of extracellular A in cerebral plaques still remains to be elucidated, increasing evidence suggests that intraneuronal A may be involved as an early event of AD pathogenesis.13,19-23 Immunohistochemical studies have demonstrated Aβ42 IR-positive neurons particularly in young DS cases. Gouras et al. observed cytoplasmic staining in neurons using antibody QCB42 in a 3-year-old DS case.23

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FIGURE 12.4 BC42 vascular immunostaining in DS brain. (a) Smooth muscle cell cytoplasmic BC42 immunoreactivity (arrowheads) in a leptomeningeal blood vessel in the temporal cortex of a 16-year-old DS patient. (b and c) Adjacent 8-µm sections of temporal cortex from a 73-year-old DS patient were immunostained with BC42 (b) and BC40 (c). Large arrowheads show colocalization of BC42 and BC40. Small arrowheads show lack of colocalization. Bars = 100 µm. (Reprinted from Lemere C.A. et al., in Neurobiology of Disease, Vol 3, 1996, p. 26. With permission from Elsevier.)

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FIGURE 12.5 Aβ42 is detected in much greater quantity than Aβ40 in PS1-FAD brains. (a) Large numbers of Aβ42-IR plaques are present in frontal cortex, including compacted plaques in all layers (small and medium arrowheads) and a diffuse Aβ42 band in layer IV (large arrowhead). (b) A few compacted Aβ40-IR plaques (small arrowheads) and blood vessels occur in an adjacent section to the one shown in a. Note that most of the Aβ40-positive blood vessels are also Aβ42-IR (asterisk). (c) Many intensely Aβ42-IR plaques (arrowhead) are present just outside the dentate gyrus and in CA1 and subiculum in the hippocampus. (d) A subset of Aβ42-containing plaques are also Aβ40-IR (for example, arrowheads in d and c) in a section adjacent to that shown in c. (e) Numerous Aβ42-IR plaques occur in cerebellum, including diffuse plaques in the molecular layer (large arrowhead) and compacted plaques in the molecular, Purkinje cell and granule cell layers (small arrowheads). Many leptomeningeal blood vessels are also Aβ42-IR (asterisk). (f) A minority of compacted plaques in the Purkinje cell layer (left arrowhead) and molecular layer (right arrowhead) and leptomeningeal blood vessels (for example, asterisk) are labeled by Aβ40 antibody in an adjacent section to that shown in e. Sections a through d are from a 47-year-old patient with an E280A PS-1 mutations; sections e and f are from a 62-year-old patient with the same PS-1 mutation. Scale bars = 500 µm. (Reprinted from Lemere C.A. et al., in Nature Medicine, Vol 2, 1996, p. 1147. With permission from Nature, http://www.nature.com/.)

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Another DS study reported TUNEL-positive neurons immunolabeled for Aβ42, indicating apoptosis in neurons containing abundant Aβ42.20 Our study also detected intraneuronal Aβ42 immunolabeling in very young (3 and 4 years old) DS cases (Figure 12.6a). However, Aβ42-immunoreactive neurons decreased to very few or none with aging and increasing plaque deposition (Figure 12.6b through Figure 12.6d).13 Intraneuronal Aβ was nearly absent in DS brains displaying dystrophic neurites, cored plaques, and neurofibrillary tangles.

β 12.3.4 INTRANEURONAL Aβ

IN

APP TRANSGENIC MICE

Intraneuronal Aβ seems to be an early step in AD pathogenesis as supported by DS and other nonDS studies. For example, using immuno-gold EM and monoclonal antibody (Mab) MBC42, Takahashi et al. reported Aβ42 localization to multivesicular bodies of neurons in Tg2576 mice with human APP Swedish 670/671 mutation.21 They speculate that Aβ42 accumulation in presynaptic and postsynaptic compartments causes synaptic dissolution. Using triple transgenic mice models with APP, PS1, and tau mutations, Oddo et al. reported impairment in synaptic plasticity correlating with intraneuronal Aβ accumulation.19 However, their study utilized Mabs 4G8 and 6E10 to detect anti-Aβ; both Mabs are known to cross-react with APP. Thus, it is possible that intraneuronal staining observed using those antibodies was APP.

12.3.5 APP IMMUNOREACTIVITY Amyloid precursor protein (APP) involves two series of endoproteolytic cleavages by secretases to produce Aβ. The first cleavage by a recently identified novel membrane-bound aspartyl protease, β-secretase (BACE1), generates an APP C-terminal fragment known as C99.24 Next, a γ-secretase complex containing PS1 and nicastrin cleaves the C99 fragment.25 Confocal microscopy demonstrated the presence of BACE1 and APP in late-Golgi.26 In our IHC studies using APP-specific antibodies, we found a vesicular staining pattern of APP in subcellular compartments or sometimes around the nuclei of neurons in human DS and AD brains and in mice overexpressing mutant APP (Figure 12.7a through Figure 12.7c). APP antibodies also recognized neuritic processes in compacted plaques in human and transgenic mouse brains (Figure 12.7b and Figure 12.7c).

12.3.6 COLOCALIZATION

OF

β Aβ

WITH

GLIA

Pro-inflammatory responses occur in pathologically vulnerable areas of the AD brain. Activated microglia and reactive astrocytes cluster within or in proximity to neuritic plaques. When activated, those inflammatory cells show ramified processes and interdigitate into the plaques. Microglia cells interacting with Aβ were observed by double immunofluorescent IHC using Pab R1282 (a general Aβ antibody) and Mab CD45 (to detect activated microglia) in PSAPP transgenic mice regardless of whether the animals had been immunized against Aβ peptide (Figure 12.8). Mounting evidence from animal studies suggests that fibrillar amyloid peptide triggers the activation of microglia and astrocytes that clearly colocalize with Aβ plaques.27,28 It seems as though inflammatory cells remove Aβ by phagocytosis, as

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FIGURE 12.6 Aβ42 staining with Pab C42 in DS subjects of increasing ages. (a) Intense cytoplasmic neuronal staining (arrows) in the absence of extracellular Aβ IR was evident at 3 years. Scale bar = 10 µm. (b) Temporal cortices of many young DS patients such as the 17year-old shown here contained both diffuse plaques (large arrowhead) and neuronal staining (small arrowhead). Scale bar = 50 µm. (c) Fully matured plaques (arrow) along with some diffuse plaques (arrowhead) were present as early as 29 years of age. Neuronal staining was very infrequent in this subject and generally seemed to decline with age as plaques matured (see Table 12.1). Scale bar = 20 µm. (d) At 62 years of age, mature cored plaques were present (arrow) and intraneuronal Aβ42 was nearly absent. Scale bar = 10 µm. (Reprinted from Mori C. et al., in Amyloid: J. Protein Folding Disorders, Vol 9, 2002, p. 95. With permission from Parthenon/CRC Press/Taylor & Francis, http://www.tandf.co.uk.)

demonstrated in vitro29 and in vivo.30 Moreover, microglia-mediated cytokines such as TNF-α and IL-6 exhibit neurotoxic as well as neuroprotective effects. The paradoxical roles of inflammatory cells in the neurodegenerative AD brain need to be clarified further.

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FIGURE 12.7 APP immunoreactivity. (a) APP Mab 8E5 detected neuronal cytoplasmic vesicles (arrows) in frontal cortex of a 52-year-old DS subject. Original magnification × 100. (b) Mab 8E5 detected cytoplasmic APP in neurons (arrows) and plaque-associated dystrophic neurites (arrowheads) in temporal cortex of a 75-year-old AD patient. Original magnification × 32. (c) Neuritic plaques (arrowheads) and intraneuronal human APP (arrows) were immunolabeled by Mab 8E5 in entorhinal cortex of an 18-month-old APP transgenic mouse overexpressing a familial mutation in human APP. Original magnification × 32.

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FIGURE 12.8 (See color insert following page 114.) Double immunofluorescent labeling of plaques (R1282) and microglia (CD45) showed similar patterns of colocalization in Aβ immunized (bottom) and untreated (top and middle) PSAPP mice after 8 weeks of Aβ immunization. A dramatic reduction in plaque burden was seen in the 13-week-old mice after treatment. CD45-immunoreactive microglia colocalized with compacted plaques in treated and untreated mice. However, because the numbers of plaques were significantly reduced in the Aβ immunized mice, the number of labeled microglia was also reduced. Images were obtained using a Zeiss Axiovert 100 M laser-scanning confocal microscope (LSM510). (Reprinted from Lemere C.A. et al., Modulating amyloid-beta levels by immunotherapy: A potential therapeutic strategy for the prevention and treatment of Alzheimer’s disease, in Saido, T.C., Ed., Amyloid-beta Metabolism and Alzheimer’s Disease, Landes Bioscience, Georgetown, TX, 2003, p. 155. With permission from Landes Bioscience, http://www.Eurekah.com.)

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REFERENCES 1. Glenner, G.G. and Wong, C.W. Alzheimer’s disease: initial report of the purification and characterization of a novel cerebrovascular amyloid protein, Biochem. Biophys. Res. Commun., 120, 885, 1984. 2. Hardy, J. Framing β-amyloid. Nature Genet., 1, 233, 1992. 3. Joachim, C.L. et al. Protein chemical and immunocytochemical studies of meningovascular β-amyloid protein in Alzheimer’s disease and normal aging. Brain Res., 474, 100, 1988. 4. Levy, E. et al. Mutation of the Alzheimer’s disease amyloid gene in hereditary cerebral hemorrhage, Dutch-type. Science, 248, 1124, 1990. 5. Masters, C.L. et al. Amyloid plaque core protein in Alzheimer disease and Down syndrome. Proc. Natl. Acad. Sci. USA, 82, 4245, 1985. 6. Mullan, M. et al. A pathogenic mutation for probable Alzheimer’s disease in the APP gene at the N-terminus of β-amyloid. Nature Genet., 1, 345, 1992. 7. Neve, R.L. et al. Expression of the Alzheimer amyloid precursor gene transcripts in the human brain. Neuron, 1, 669, 1988. 8. Selkoe, D.J. et al. Isolation of low-molecular-weight proteins from amyloid plaque fibers in Alzheimer’s disease. J. Neurochem., 146, 1820, 1986. 9. van Broeckhoven, C. et al. Amyloid β-protein precursor gene and hereditary cerebral hemorrhage with amyloidosis (Dutch). Science, 248, 1120, 1990. 10. Wisniewski, K.E., Wisniewski, H.M., and Wen, G.Y. Occurrence of neuropathological changes and dementia of Alzheimer’s disease in Down’s syndrome. Ann Neurol., 17, 278, 1985. 11. Mann, D.M. et al. A morphological analysis of senile plaques in the brains of nondemented persons of different ages using silver, immunocytochemical and lectin histochemical staining techniques. Neuropathol. Appl. Neurobiol., 16, 17, 1990. 12. Lemere, C.A. et al. Sequence of deposition of heterogeneous amyloid beta-peptides and APOE in Down’s syndrome: implications for initial events in amyloid plaque formation. Neurobiol. Dis., 3, 16, 1996. 13. Mori, C. et al. Intraneuronal Aβ42 accumulation in Down’s syndrome brain. Amyloid: J. Protein Folding Disord., 9, 88, 2002. 14. Iwatsubo, T. et al. Amyloid beta protein (Aβ) deposition: A beta 42(43) precedes A beta 40 in Down’s syndrome. Ann. Neurol., 37, 294, 1995. 15. Joachim, C.L. et al. Protein chemical and immunocytochemical studies of meningovascular β-amyloid protein in Alzheimer’s disease and normal aging. Brain Res., 474, 100, 1988. 16. Jarret, J.T., Berger, E.P., and Lansbury, P.T., Jr. The carboxy terminus of the beta amyloid protein is critical for the seeding of amyloid formation: implications for the pathogenesis of Alzheimer’s disease. Biochemistry, 32, 4693, 1993. 17. Lemere, C.A. et al. The E280A presenilin 1 Alzheimer mutation produces increased Abeta 42 deposition and severe cerebellar pathology. Nat Med., 2, 1146, 1996. 18. Miklossy, J. et al. Two novel presenilin-1 mutations (Y256S and Q222H) are associated with early-onset Alzheimer’s disease. Neurobiol. Aging, 24, 655, 2003. 19. Oddo, S. et al. Triple-transgenic model of Alzheimer’s disease with plaques and tangles: intracellular Abeta and synaptic dysfunction. Neuron, 39, 409, 2003. 20. Busciglio, J et al. Altered metabolism of the amyloid β precursor protein is associated with mitochondrial dysfunction in Down’s syndrome. Neuron, 33, 677, 2002. 21. Takahashi, R.H. et al. Intraneuronal Alzheimer Aβ42 accumulates in multivesicular bodies and is associated with synaptic pathology. Am. J. Pathol., 161, 1869, 2002.

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22. Gyure, K.A. et al. Intraneuronal Aβ-amyloid precedes development of amyloid plaques in Down syndrome. Arch. Pathol. Lab. Med., 125, 489, 2001. 23. Gouras G.K. et al. Intraneuronal Aβ42 accumulation in human brain. Am. J. Pathol., 156, 15, 2000. 24. Vassar, R. et al. Beta-secretase cleavage of Alzheimer’s amyloid precursor protein by the transmembrane aspartic protease BACE. Science, 286, 735, 1999. 25. Yu, G. et al. Nicastrin modulates presenilin-mediated notch/glp-1 signal transduction and beta APP processing. Nature, 407, 48, 2000. 26. Yan, R. et al. The transmembrane domain of the Alzheimer’s beta-secretase (BACE1) determines its late Golgi localization and access to beta-amyloid precursor protein (APP) substrate. J. Biol. Chem., 276, 39, 36788, 2001. 27. Benzing, W.C. et al. Evidence for glial-mediated inflammation in aged APP(SW) transgenic mice. Neurobiol. Aging, 20, 581, 1999. 28. Schenk, D. et al. Immunization with amyloid-beta attenuates Alzheimer disease-like pathology in the PDAPP mouse. Nature, 400, 173, 1999. 29. Paresce, D.M., Ghosh, R.N., and Maxfield, F.R. Microglial cells internalize aggregates of the Alzheimer’s disease amyloid beta-protein via a scavenger receptor. Neuron, 17, 553, 1996. 30. Bard, F. et al. Peripherally administered antibodies against amyloid beta-peptide enter the central nervous system and reduce pathology in a mouse model of Alzheimer’s disease. Nat. Med., 6, 916, 2000.

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13

Creating APP Transgenic Lines in Mice Stanley Jones Premkumar Iyadurai and Karen Hsiao Ashe

CONTENTS 13.1 Introduction 13.1.1 Origins of Gene Manipulation and Gene Transfer into Mouse Genome 13.1.2 Basic Principles for Consideration in Generating Transgenic Mice 13.1.3 APP Gene Structure: Its Isoforms and Promoters Used in Creating APP Transgenics 13.2 Experimental Procedures 13.2.1 Steps in Generating APP Transgenic Mice by Microinjection 13.2.1.1 Strain Selection 13.2.1.2 Isolation of Target DNA from Bacterial Host 13.2.1.3 Purification of DNA for Microinjection 13.2.1.4 Preparation of DNA for Microinjection 13.2.1.5 Microinjection with Admixed DNAs 13.2.1.6 Indentification of Founders 13.2.1.7 Maintenance and Analysis of Founders 13.3 Concluding Remarks References

13.1 INTRODUCTION Alzheimer’s disease (AD) is a progressive neurodegenerative disease that primarily impairs memory function.1 Pathologically, AD is characterized by amyloid-containing neuritic plaques and intraneuronal fibrillary tangles.2 While the biological causes and exact biological mechanisms by which the progressive neurodegeneration is initiated are not very clear, it has been shown that mutations in some specific genes may be important in these events. For example, mutations in genes encoding the amyloid precursor protein (APP),3–9 presenilin-1 (PS1) and presenilin-2 (PS2)10–13 have been associated with cases of familial AD.

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Further analyses have shown that mutations in all the aforesaid genes result in increased amounts of a specific cleavage product of APP, namely, Aβ.14–17 Interestingly, Aβ has been identified as a component of neuritic plaques in AD brains18 and lends support to Aβ serving as a key modulator in the progression of AD. Another gene encoding the apolipoprotein E (apo E) is also thought to be a modifier of the Alzheimer’s disease phenotype.19 Although AD was first described almost a century ago,20 the sequential events that lead to AD and memory dysfunction are not clearly defined and remain hard to elucidate for several reasons. At present, AD is a clinical diagnosis; definitive diagnosis is available only upon autopsy. Because of the variations in presentation and overlapping disease symptoms with other neurological disorders, AD poses a challenge for clinical diagnosis. For moral and ethical reasons, antemortem biopsy of brain tissue for diagnosis is not a viable option and studying the pathogenesis in humans has been very difficult. However, the identification of genes associated with AD and the advent of molecular biological tools for gene transfer have made it possible for AD researchers to study the human AD-causing genes in vivo in basic model organisms such as mice, rats, flies, worms, and yeasts. One approach is to study the homologous genes in a given model system, for example, studying the mouse homologue of the human PS gene in mice. Another approach is to transfer and express the human gene of interest in a model organism, for example, expressing the human APP gene and studying the consequences of expressing it in mice. The latter technique has been termed the transgenic mouse approach for modeling human diseases.

13.1.1 ORIGINS OF GENE MANIPULATION INTO MOUSE GENOME

AND

GENE TRANSFER

The laboratory mouse remains the well-studied genetic model organism that is evolutionarily closest to the human. With the rediscovery of Mendel’s laws in the 1900s and subsequent interest in experimental mouse developmental biology, the field of mouse genetics was firmly established by the 1960s.21 The first report of introduction of foreign DNA into a transgenic mouse as a way of manipulating the mouse genome emerged from Jaenisch and Mintz in 1974.22 These researchers showed that purified SV40 DNA, when injected into the blastocoel cavities of mouse blastocysts, was capable of integrating into the genomes of the embryonic cells. Following the discovery that microinjection of cloned herpes simplex virus thymidine kinase (tk) DNA into the nuclei of cultured fibroblasts led to stable expression of the tk gene,23,24 Gordon et al.25 reported the first successful pronuclear injection into a one-celled mouse embryo and demonstrated that the transgenic mouse showed expression in somatic tissues. Integration of such foreign DNA into somatic and germline tissues was reported immediately afterward.26 Since then, a huge number of genes, including human genes, have been introduced into the mouse genome. The establishment of human disease conditions in mice revolutionized the research arena, especially in understanding complex diseases such as AD. While no

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genetic disease model in mice can supplant the need for studying the disease in humans and human clinical trials, studies in model systems allow researchers to gain some understanding of the pathology and coordinate efforts to approach the disease in humans. The ability to transfer human genes to mice raised the possibility of studying mutant disease-causing human genes in the context of the mouse genome and establishing model disease paradigms. Research has shown that mouse disease model systems, albeit partial model systems, mimic human disease processes and serve as valuable tools for understanding pathogenesis, progression, prevention, and treatment. Several human disease models exist in transgenic mice, including AD,27 Parkinson’s disease,28 Huntington’s disease,29,30 amyotropic lateral sclerosis,31 and others.32,33 The key first step in developing a model system is to identify specific genes involved in the disease condition. This step is usually carried out by following family pedigrees manifesting a given disease and using molecular biological approaches. Once a gene is identified and cloned, one can study the basic biologic functions of the gene in heterologous systems, such as mice, by expressing that human gene in mice. For example, in a familial form of early-onset AD observed in Sweden, the offending gene mutation was identified in the amyloid protein precursor (APP) gene.3 The cDNA expressing the “Swedish mutation” of the APP gene was later cloned and transferred to mice.34 The principles and methodology of generating transgenic mice will be dealt in detail in the sections to follow. In addition to the transgenic technology, two other methodologies have been used to manipulate the mouse genome — knock-out and knock-in mice — to understand the pathogenetic mechanisms underlying human diseases. Both technologies utilize the properties of homologous recombination and embryonic stem (ES) cells. ES cells are derived from cells of the inner cell masses in developing blastocysts early in embryonic development.21 ES cells can be isolated and cultured35,36 and manipulated to include foreign genes in culture.37 Such manipulated ES cells can be reinjected into a developing mouse blastocyst and then implanted into a recipient. If the manipulated ES cells go on to differentiate into the germline, among other tissues, a germline transgenic mouse is generated. The knock-out strategy involves replacement of the wild-type gene with a disrupted version of a given gene (usually with neoR) or with a truncated version of the gene, so that the resulting gene at its native location is nonfunctional or dysfunctional. The knock-in strategy involves replacement of the wild-type gene, with a specifically engineered mutation of the same gene. The advantage of generating knock-in and knock-out models is that the given gene is expressed from its native location in its own transcriptional context, usually mimicking the native gene’s expression abundance levels. Patterns and abundance expression at endogenous levels have often been insufficient to produce desirable phenotypes. Moreover, since both knock-in and knock-out techniques rely on extensive homology of DNA sequences for homologous recombination, replacing native mouse genes with mutant human genes is limited to genes with high degrees of DNA homology. The generation of transgenic animals by microinjection overcomes these limitations and thus remains a superior strategy compared to other strategies discussed.

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13.1.2 BASIC PRINCIPLES FOR CONSIDERATION TRANSGENIC MICE

IN

GENERATING

An ideal disease model in any model organism recreates the typical features of the human disease and its progression. To this end, effective design of the transgene, its expression profile and strength of expression are very important factors. Therefore special consideration should be given to the transgene construction — whether to use the native gene or the complete cDNA and the relevant choice of promoters for favorable expression — with expected expression profiles in a spatial and a temporal fashion, and with expression strong enough to recreate the mutant phenotype. In the case of cDNA-based transgenic mice, the choice of one isoform among others derived from the same gene is critical. While creating the transgenic mouse with the full native gene circumvents the problem of isoform selection, selective cDNA expression may be advantageous when the given cDNA is more closely related to the development of the disease, for example, an isoform with neuron-specific expression (as opposed to ubiquitous expression) to study a neurological disorder. Additionally, one can use specific promoters to drive expression in a certain subset of cell types in mice. It should also be borne in mind that since transgenic mice are generated by insertion of the gene of interest at a random site in the mouse genome, the gene is influenced by “position effect” — the expression of the gene is influenced by the location where it is integrated.38 For the same reason, it should be expected that the expression levels in independent germline transgenic mice carrying the same transgene will be different. Based on this consideration, it is essential to generate multiple, independent transgenic lines to ensure that one of them will provide sufficient expression to produce the mutant phenotype. Traditionally, several types of promoters have been used in directing expression of transgenes in transgenic mouse lines. The promoters derived from the so-called “housekeeping” genes have been known to direct ubiquitous expression. These promoters include the β-actin promoter,39,40 the mouse metallothionein promoter,41,42 the HMG CoA reductase promoter,43 and the histone H4 promoter.44 It should be noted that although the promoters are “ubiquitous,” they may not direct the same levels of expression in all tissues. Another class of promoters, the conditional promoters, has also been used successfully. In their case, the transgene of interest is silent until it is activated by specific manipulation, such as heat shock or by an inducer. Specific examples of this class of promoters include metallothionein promoter (inducers: Zn and Cd), hsp68 promoter (inducer: heat shock), lac operatorpromoter (inducer: IPTG), and tet operon promoter (inducer: tetracycline). Other strategies for conditional expression involving FLP sites and FLP recombinase45 loxP sites and P1 Cre recombinase46 are beyond the scope of this chapter.

13.1.3 APP GENE STRUCTURE: ITS ISOFORMS AND PROMOTERS USED IN CREATING APP TRANSGENICS APP, whose function is not completely clear, is the precursor of Aβ which is a core component of senile plaques in AD brains.47 The APP gene contains 19 exons and is known to undergo alternative splicing.48 APP is known to have at least five major

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isoforms of sizes 639,49 695, 714, 751, and 770 amino acids.47,50 The longest isoform, APP770, contains a 56-amino acid domain (encoded by exon 7), which shares sequence homology with and can function like Kunitz-type serine protease inhibitors (KPIs)51 as well as an adjacent 19-amino acid domain (encoded by exon 8) with homology to the MRC OX-2 antigen1,52 found on the surfaces of neurons and certain immune cells. 53 Brain tissue expresses little APP770. APP751 contains the KPI region but lacks the MRC OX-2 domain. APP751 is expressed variably in the brain at low, intermediate, or high levels, depending on the region involved. APP695 lacks both of the above domains and is primarily produced in neurons that constitute the primary sources of APP in the central nervous system. APP714 contains amino acids of APP695 form plus an addition of 19 amino acids of unknown significance, and is not present in the brain. APP639 also lacks exon 2 and is highly expressed in liver and least expressed in the brain.49 Several transgenic AD models have been generated using expression of modified APP, PS1, and tau genes. Most groups have focused on generating transgenic mouse lines carrying mutant APP or PS1 genes. In this section, the discussion will be limited to selected APP transgenics. The first APP transgenic model of AD in mice was created by expressing human wild-type APP751 driven by a neuron-specific enolase promoter in inbred JU mice that developed age-related impairments in memory tests.54,55 Although no amyloid plaques were reported, diffuse Aβ/APP deposits and abnormal tau immunoreactivity were observed in aged animals.56 The second APP transgenic model of AD in mice was created by Games and colleagues.57 These authors expressed a recombinant transgene derived from portions of APP cDNA and genomic DNA fragments such that alternative splicing of exons 7 and 8 could be functional. The recombinant transgene was driven by the plateletderived growth factor β-chain promoter.57 In addition, the transgene contained V717F, the “Indiana” mutation. These mice exhibited amyloid deposition around 6 to 9 months, but neurofibrillary tangles were absent.58,59 Tg2576 was generated by injecting APP cDNA containing the “Swedish mutation” (K670N, M671L) under the control of the hamster prion protein promoter.59 The prion protein promoter drives expression pan-neuronally in the brain.60 Plaque deposition and cognitive dysfunction in an age-dependent fashion have been described.61 Another set of transgenic lines, TgAPP22 and TgAPP23, were generated using APP751 cDNA containing the Swedish mutation, under the neuron-specific Thy-1 promoter.62 Thy-1 promoter also directs expression throughout the central nervous system. TgCRND8 mice were generated by expressing APP695 cDNA containing both the Swedish and Indiana mutations under the control of the pan-neuronal hamster prion protein promoter.63 These mice developed memory deficits at the early age of 3 months. APP knock-in or yeast artificial chromosome-based mice have also been generated and they tend to develop pathologies later in life.64 The utility of transgenic and knock-out mice in modeling neurological diseases is outlined in Aguzzi et al.65 The knock-in models have more recent origins, relatively speaking. Most of the knock-in models of AD have been described in the presenilin

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genes. Mutations in PS1 and PS2 have been linked to familial AD. The first ADmodeling knock-in mice were reported by Guo and colleagues.99 These researchers exchanged the wild-type PS1 sequences for a mutant PS1 sequence to make a methionine-to-valine change at the 146th amino acid position.66 Although the mice did not exhibit any overt phenotype, they were hypersensitive to seizure-induced synaptic degeneration and necrotic neuronal death in the hippocampus. Another PS1 knock-in (PS1P264L), using a similar strategy was reported by Siman et al.67 Apolipoproteins E4 and E3 are alleles of apo E that seem to modify AD pathology, although the precise function of apo E in the AD pathway is unknown. To understand the roles of apo E4 and apo E3 mutations in cholestrol metabolism in synaptic plasma membranes, knock-in models of apo E4 and E3 were generated by replacing the wild-type apo E by homologous recombination.68 These authors found that the apo E4 knock-in mice showed abnormal cholestrol distribution in the synaptic plasma membrane, as compared to wild-type and Apo E3 knock-in mice and suggested that the pathogenic effects of apo E4 might be mediated by the altered synaptic plasma membrane. To better understand the role of APP in development, native mouse APP was deleted from the mouse genome by homologous recombination.69 The resultant “APP null” mice were viable and fertile, showing mild deficits in locomotor activity and reactive gliosis, indicating a role for APP in normal neuronal function. Efforts have been successful in developing an AD model system containing modified APP transgenes that can be induced to express at will in mice. We have used the tetracycline activator–tetracycline-response element (TRE) system to express APP and tau transgenes in our laboratory. In brief, constructs containing modified APP and tau transgenes were cloned behind the TRE. The constructs were then microinjected into pronuclei of one-celled embryos and transgenic mice were produced. These mice were the transgenic responders. The activator mice were created by Mayford et al.70 (a kind gift of Dr. Eric Kandel, Columbia University, New York). The mice expressed the tetracycline transactivator (tTA) in the forebrain, as dictated by the calcium–calmodulin-dependent kinase II (CaMKII) promoter regulatory elements, behind which the tTA was cloned (CKII-tTA).70 The CKII-tTA fusion ensures that the tTA expression mimics the native CKII expression profile in a spatial- and temporal-specific fashion. Under normal circumstances, the responder gene is silent. However, in mice that are doubly transgenic for CKII-tTA and TRE-responder (APP or tau), the APP or tau expression, as appropriate, was observed. As expected, the CKII-tTA/TRE responder-mediated induced gene expression in the double transgenic mice could be eliminated by addition of tetracycline to the diet or water. Thus, in the CKII-tTA/TRE responder double transgenics, in the absence of tetracycline, the responder gene was expressed; in the presence of tetracycline, the responder gene was repressed. Models such as these will be effective in answering a variety of questions related to time-specific expression of modified APP and tau transgenes and the consequences. The quest for creating a “complete” model for AD is ongoing. Ideally, such a model would develop senile plaques and neurofibrillary tangles in a time-dependent fashion and show time-dependent cognitive decline, similar to conditions observed

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in humans. Such a model does not exist. However, attempts have been made to combine available transgenic lines containing mutations in APP, PS1 and tau genes by conventional breeding. Some of these lines are very promising. In the course of breeding mice carrying the tauP301L mutation and Tg2576, we have shown that mice doubly transgenic for tauP301L and APPK670N, M671L exhibit amyloid plaques and neurofibrillary tangles (data not shown). Bringing transgenes together by conventional breeding is limited by lack of maintenable homozygosity (due to insertional mutations from the transgene), strain effects on the transgene, etc. An ingenious approach to creating a closer-to-real model was taken by Oddo et al.71 They first generated a PS1 knock-in mutant transgenic mouse (PS1M146V) and then microinjected APPK670N, M671L and tauP301L transgenes into single-cell embryos derived from the homozygous PS1M146V mice. They were able to show that amyloid deposition precedes tangle formation in this triple transgenic model.72 In the yet-tobe developed “ultimate model,” tau pathology should be generated from wild-type tau since mutations in tau are not associated with AD.73

13.2 EXPERIMENTAL PROCEDURES 13.2.1 STEPS

IN

GENERATING APP TRANSGENIC MICE

BY

MICROINJECTION

13.2.1.1 Strain Selection It is conceivable that the expression of a transgene and its behavior in one strain background may be different from the expression in another strain because different host strains contain different modifier alleles that may interact with the transgene or the gene at the point of insertion. This effect is true in the case of mice expressing APP transgenes as well. Our laboratory generated Tg(HuAPP695.SWE)2576 mice by microinjecting C57B6j × SJL F2 eggs.60 The Tg2576 transgene array could not be transferred onto the C57B6j inbred background because the proportion of mice dying prematurely increased and the fraction of transgene-positive mice that were weaned fell significantly below the expected 50% as the percentage of C57B6jderived alleles increased. 74 Interestingly, when these mice were crossed with F1 hybrids of C57B6j x SJL, the fraction of mice living long term increased, indicating that the SJL-derived alleles protected against the lethal effects of APP overexpression. Similarly, when APP transgenes were expressed in FVB/N mice, premature death was usually preceded by a variety of neurologic signs including neophobia and thigmotaxic behavior.34 It was later shown that the influence of FVB/N-derived alleles was enough to cause behavioral abnormalities when a transgene generated in the C57B6j x SJL mice was crossed to FVB/N mice.60 Concentrations of APP that produce amyloid plaques in outbred transgenic lines were lethal for inbred FVB/N or C57B6j mice. From our experience, we chose to create transgenic mice that overexpress APP by microinjecting C57B6j x SJL F2 embryos. It should also be noted that because of the relatively poor reproductive performance of inbred mice, the production of fertilized eggs and the generation of transgenic mice and their subsequent breeding are more efficient when F2 zygotes are used for microinjection.75 However, we have successfully used FVB/N mice for generation of TRE

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responder transgenic mice. FVB/N mice offer the advantages of being an inbred strain with relatively large sized pronuclei in fertilized eggs, being fairly resistant to lysis during injection and exhibiting good reproductive characteristics.76 13.2.1.2 Isolation of Target DNA from Bacterial Host Isolation of DNA for the purposes of microinjection is a critical step. Usually the source of the DNA is plasmid DNA and occasionally, cosmid DNA. Since the dissolved DNA is injected ultimately into a viable zygote, the DNA should not contain any chemical or compound that will be toxic to the early zygote and its development. First, it is essential to grow the bacteria harboring the plasmid or the cosmid in the right host. We found that the use of Escherichia coli host strains DH5α and XL-1 Blue produce similar results. However, it should be borne in mind that if special host factors are required for maintenance of the plasmid or cosmid, it may be appropriate to use a special, specific host. We routinely harvest bacteria after 15 to 16 hours of growth at 37ºC. Storing bacterial pellets at –20ºC does not seem to affect the quality of the isolated DNA. The purity of DNA and the solution in which the DNA is suspended are of paramount importance. For this reason, traditional DNA purification methodologies involving cesium chloride gradients or ethidium bromide are not usually performed. Subjecting the DNA to restriction enzyme digestion and DNA sequencing analysis can readily determine purity of the DNA samples. Pure DNA that is free of proteins and other contaminants will be readily cut with restriction enzymes and may be readily sequenced with standard methods. Our laboratory has successfully used commercially made kits (QIAGEN; http:www.qiagen.com) that employ modified alkaline lysis procedures to isolate pure DNA. In brief, bacterial cell lysis is achieved, followed by binding the DNA to a column and eluting the DNA. The quality of DNA can be cross-checked by effective restriction analysis and/or the amenability of the DNA to sequencing. In addition, in our laboratory, we have obtained better results with kits that yield endotoxin-free DNA. In protocols for endotoxin-free DNA, the additional step of entoxin removal must be performed before the DNA is bound to the column for elution. What is the biggest construct size one can use to generate transgenic mice? The answer: as big as anyone can make it. Precedents in the literature document stable integration of a 50-kb bacteriophage λ clone,77 a 60-kb cosmid insert,78 and a 70-kb fragment produced by in vitro ligation of two cosmid inserts.79 As in yeasts, coinjection of large sequences with overlapping homologous regions allows homologous recombination to effectively reconstruct the linear sequence.80 In addition, yeast artificial chromosomes that contain 300 to 400 kb of DNA have also been successfully injected to produce transgenic mice by pronuclear injection.81 We successfully generated several lines of transgenic mice by injecting cosmid DNAs up to 50 kb in size. Whether introns are necessary for favorable expression of the desired gene product is not clear. It has been shown that the levels of gene expression from cDNAbased constructs are significantly lower than those obtained with genomic sequences,

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which includes introns and exons.82 It has been suggested that enhancers in the intronic sequences may be one reason for this observation. However, it has also been noted that the addition of heterologous introns to cDNA-based constructs can yield increases in gene expression levels.44,83 13.2.1.3 Purification of DNA for Microinjection For microinjection into fertilized oocytes, plasmid or cosmid DNAs are further processed to generate specific DNA fragments without any native bacterial DNA vector sequences. Although prokaryotic cloning vector sequences have no apparent effects on the integration frequency of microinjected genes, it is important to note that they can severely inhibit the expression of eukaryotic genes introduced into the mouse germline.84–87 Brinster et al.75 showed that both linear and supercoiled DNAs are capable of integration into the mouse genome; however, linear DNA integrated at a greater frequency (25% vs. 5%). The specific processing goals include (1) using specific restriction endonucleases that strictly flank the DNA of interest, (2) separating the DNA fragments on a gel, (3) eluting the DNA fragment from the gel, and (4) reconcentrating the DNA fragment if necessary. Care should be taken to make sure that the DNA fragment of interest does not contain any site for the specific restriction endonuclease used to remove the bacterial vector sequences, even within the intronic regions. Restriction digests are performed per standard conditions; however, the conditions of electrophoresis are modified so that the DNA is not contaminated with ethidium bromide used in standard gel separation methodologies. In fact, several modifications are employed in the electrophoresis protocol. First, the appropriate percentage agarose gel is cast without included ethidium bromide. The gel contains a slab well and at least two normal wells. The bulk of the restriction enzyme-cut DNA is loaded to the slab well and a small-volume sample is loaded next to a lane of standard molecular weight markers. After appropriate hours of running the gel, the marker and the small-volume sample lanes are cut and separately stained with ethidium bromide. Once the bands are visualized, they are aligned to the original gel and the DNA fragment of interest is cut out. The cut-out DNA fragment still embedded in the gel is placed into treated dialysis tubing containing 0.1X TAE. The DNA fragment is eluted from the agarose gel onto the 0.1X TAE by placing the tubing inside an electrical field, usually the gel electrophoresis chamber. For a starting amount of 20 µg DNA, the gel is run for 2 hr. The eluted DNA remains in solution inside the dialysis bag and is then transferred to a fresh Eppendorf tube. If the eluted volume is too high (greater than 2 ml), pure butanol can be used to reduce the aqueous volume. The eluted DNA in solution is then precipitated by standard techniques using potassium acetate and resuspended in 5 mM Tris, 0.1 mM EDTA, pH 7.4. The concentration of the eluted DNA can be quantitatively estimated by measuring the optical density of the resultant solution at 260 nm and 280 nm. Alternatively, comparison of band intensities to similar-sized λ-Hind III fragments of known quantity can be done to estimate the quantity of the eluted DNA fragment.

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13.2.1.4 Preparation of DNA for Microinjection DNA samples for microinjection should be free of contaminants such as traces of phenol, ethanol or other proteinaceous debris. When the concentration of the DNA solution containing the fragment of interest is established, the DNA is diluted to a concentration of 4 µg/ml in 5 mM Tris, 0.1 mM EDTA, pH 7.4. For smaller constructs (<10 kb), we use a concentration of 2 µg/ml. After dilution, we have found it useful to centrifuge the solution in a table-top device at full speed (13,000 rpm) for 30 min. This approach helps prevent clogged needles during injection. If necessary, the DNA sample may also be run through a 0.2-µm filter. We have also found it useful to ensure that the DNA to be microinjected is fresh — injected on the day of reconstitution. Such DNA can be injected without problems for at least a week in the microinjection facility as long as it is stored at 4ºC. In general, concentrations above 1 µg/ml75 or 1 to 3 µg/ml of linear DNA yield a DNA integration efficiency of 20 to 40%. We have obtained an integration efficiency of about 20%. 13.2.1.5 Microinjection with Admixed DNAs It is known that DNA injected into the nucleus is incorporated into the native mouse genome. Therefore, in theory, two distinct DNA fragments, if injected, should also be incorporated into the mouse genome, presumably independent of each other. It was reported earlier that two genes mixed and coinjected into mouse eggs generated transgenic mice carrying both transgenes.88 It was also noted that both transgenes cointegrated at the same site in the genome in a head-to-tail fashion. We successfully used this principle to generate a mouse containing both APP (Swedish and London [V717I] mutations) and tauP301L mutations. In brief, DNA fragments representing modified APP and modified tau transgenes were coinjected into fertilized mouse oocytes. Few of the transgenic mice generated from these coinjection studies contained both transgenes. Some had one or the other transgene, as confirmed by Southern blot hybridization (data not shown). By careful breeding and analysis, we have been able to separate these sublines and maintain them. We were also able to show that in mice containing both transgenes, both transgenes were expressed, as observed by Western blot analysis using antibodies against both APP and tau (data not shown). Successful recovery of multiple transgene mice using this technique raises the possibility that it may be a faster method for generating multiple transgene mice instead of the conventional method of developing individual lines of mice and breeding them. However, the integration sites of the individual transgenes may be different and the expression levels from these constructs may be different. Given this caveat, gene dosage studies using double transgenics developed in this way may be of lesser value than those developed the conventional way. Another clever methodology involving knock-in and coinjection strategies to produce triple transgenic mice containing altered APP, PS1, and tau genes was discussed earlier.71,72 13.2.1.6 Indentification of Founders Following microinjection, fertilized embryos are implanted into surrogate dams and allowed to deliver. It should be noted that the integration of injected foreign DNA

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into mouse nuclear DNA is a chance event and usually occurs about 10 to 20% of the time. Therefore, it is essential to identify the mice that are products of transgenic embryos. Polymerase chain reaction (PCR) with specific primer sets for the transgene is a sensitive enough test to screen for founders in the progeny. We routinely use PCR as the first step screen using specific primers that yield amplification products ranging between 200 and 1000 base pairs in size. To control for genomic DNA quality and amplification conditions, we also amplify a region of the mouse prion protein gene using specific primers in the same PCR reaction mixture. PCR-positive founders are subjected to maintenance breeding and analysis as warranted. PCR is a very sensitive test and utmost care should be taken while preparing the DNA and reagents and setting up the reactions. Standard precautions to avoid cross-contamination should be used. It is useful to include a reaction without DNA as a negative control every time a PCR screening is performed. PCR-positive founders are further subjected to slot blot hybridization (to rule out false positives from the PCR screen) and Southern blot hybridization (to determine the integrity of the inserted transgene). We routinely perform copy number estimation by slot blot hybridization on DNA derived from progeny of founders. This step is necessary since it is possible that the founder may have more than one insertion point, and might carry more than one array of transgene in those locations. If indeed this is the case, the progeny will show bands of varied intensities on a slot blot or bands of differing sizes on a Southern blot, implying segregation of variable copy number transgenes in multiple locations. We occasionally encountered situations where the transgene inserted in up to three locations and serial breeding to isolate stable sublines was necessary. 13.2.1.7 Maintenance and Analysis of Founders After a founder is established, it is mated with mice of the opposite sex (of the desired strain and background) for maintenance breeding. We usually combine two or three females with one male for mating. The mice are allowed for mate for 1 or 2 weeks. The pregnant females are separately housed and allowed to give birth. The litter is weaned at about 3 weeks of age, separated according to sex and housed appropriately. At weaning, the distal portions of their tails are snipped and collected to provide material for DNA extraction for analysis. DNA collected from all the progeny is subjected to PCR screening with specific primers to identify transgenic and the nontransgenic mice. DNA from mice to be used for specific experiments including behavioral, RNA, and protein analysis is further subjected to slot blot hybridization to ensure the presence or absence of the transgene. This step is necessary because PCR is so sensitive that false positives must be ruled out. Specific sublines identified and isolated from multiple-insert founders are typically followed by Southern blot hybridization with appropriate probes after appropriate restriction enzyme digestion and electrophoresis. Specific consideration should be given to experiments involving behavioral analysis where large balanced groups of transgenic and nontransgenic animals are

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required. Specific matings should ensure that approximately equal numbers of sexand age-matched transgenic and nontransgenic animals are born. While planning for behavioral experiments and calculating numbers of animals required, it is necessary to provide concessions for unexpected smaller litter size, lethality, and infertility. In the case of mice that carry regulatable transgenes, the activator line (CK-tTA mice) and the responder line (various transgenes downstream of TRE) are maintained separately and the transgene status is followed by performing PCR on DNA samples from these mice. When necessary, these lines are bred to each other to generate “double transgenic” mice. Ultimately, the transgene status of each mouse involved in the study is confirmed by slot blot hybridization.

13.3 CONCLUDING REMARKS A concerted, multidisciplinary, multifaceted approach involving genetics, molecular pathology, cell biology, biochemistry, neurophysiology, and pharmacology is necessary to understand complex diseases such as AD. The ability to create transgenic mouse models of AD has become a powerful tool for researchers aiming to understand the pathogenesis, development, prevention and treatment of AD. Several transgenic AD models with specific characteristics exist and several newer models are being generated via novel methodologies. The ability to breed transgenic AD mice that develop amyloid plaques, neurofibrillary tangles and age-dependent cognitive decline will undoubtedly allow researchers to test strategies to prevent amyloid plaque and neurofibrillary tangle formation and ultimately identify agents that prevent, delay or treat cognitive decline. Transgenic AD mice can also be useful as traditional genetic screens to identify genes that may be involved in modulating the manifestations of AD. Transgenic mice expressing APP and other genes related to AD will enable scientists to pose questions regarding the pathogenesis of AD and test relevant hypotheses relating to the memory dysfunction associated with AD. Exciting years lie ahead, when the sweet fruits of the hard labor involved in generating these transgenic AD mice will be realized.

REFERENCES 1. Muller-Hill, B. and Beyreuther, K. Molecular biology of Alzheimer’s disease, Annu. Rev. Biochem., 58, 287, 1989. 2. Wisniewski, K.E., Wisniewski, H.M., and Wen, G.Y. Occurrence of neuropathological changes and dementia of Alzheimer’s disease in Down’s syndrome, Ann. Neurol., 17, 278, 1985. 3. Mullan, M. et al. A pathogenic mutation for probable Alzheimer’s disease in the APP gene at the N-terminus of beta-amyloid, Nat. Genet., 1, 345, 1992. 4. Mullan, M. A genetic defect causing Alzheimer’s disease, Br. J. Hosp. Med., 45, 131, 1991. 5. Ancolio, K. et al. Unusual phenotypic alteration of beta amyloid precursor protein (betaAPP) maturation by a new Val-715 Met betaAPP-770 mutation responsible for probable early-onset Alzheimer’s disease, Proc. Natl. Acad. Sci. USA, 96, 4119, 1999.

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6. Chartier-Harlin, M.C. et al. Early-onset Alzheimer’s disease caused by mutations at codon 717 of the beta-amyloid precursor protein gene, Nature, 353, 844, 1991. 7. Goate, A. et al. Segregation of a missense mutation in the amyloid precursor protein gene with familial Alzheimer’s disease, Nature, 349, 704, 1991. 8. Hendriks, L. et al. Presenile dementia and cerebral haemorrhage linked to a mutation at codon 692 of the beta-amyloid precursor protein gene, Nat. Genet., 1, 218, 1992. 9. Murrell, J. et al. A mutation in the amyloid precursor protein associated with hereditary Alzheimer’s disease, Science, 254, 97, 1991. 10. Levy-Lahad, E. et al. Candidate gene for the chromosome 1 familial Alzheimer’s disease locus, Science, 269, 973, 1995. 11. Rogaev, E.I. et al. Familial Alzheimer’s disease in kindreds with missense mutations in a gene on chromosome 1 related to the Alzheimer’s disease type 3 gene, Nature, 376, 775, 1995. 12. Sherrington, R. et al. Alzheimer’s disease associated with mutations in presenilin 2 is rare and variably penetrant, Hum. Mol. Genet, 5, 985, 1996. 13. Sherrington, R. et al. Cloning of a gene bearing missense mutations in early-onset familial Alzheimer’s disease, Nature, 375, 754, 1995. 14. Cai, X.D., Golde, T.E., and Younkin, S.G. Release of excess amyloid beta protein from a mutant amyloid beta protein precursor, Science, 259, 514, 1993. 15. Citron, M. et al. Mutation of the beta-amyloid precursor protein in familial Alzheimer’s disease increases beta-protein production, Nature, 360, 672, 1992. 16. Haass, C. et al. Mutations associated with a locus for familial Alzheimer’s disease result in alternative processing of amyloid beta-protein precursor, J. Biol. Chem., 269, 17741, 1994. 17. Suzuki, N. et al. An increased percentage of long amyloid beta protein secreted by familial amyloid beta protein precursor (beta APP717) mutants, Science, 264, 1336, 1994. 18. Glenner, G.G. and Wong, C.W. Alzheimer’s disease: initial report of the purification and characterization of a novel cerebrovascular amyloid protein, Biochem. Biophys. Res. Commun., 120, 885, 1984. 19. Albert, M.S. Cognitive and neurobiologic markers of early Alzheimer disease, Proc. Natl. Acad. Sci. USA, 93, 13547, 1996. 20. Alzheimer, A. Uber eine eigenartige Erkrankung der Hirnrinde, Allg. Z. Psychiatr. Psych. Gerichtl. Med., 64, 146, 1907. 21. Hogan, B. et al. Manipulating the Mouse Embryo: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, New York, 1994. 22. Jaenisch, R. and Mintz, B. Simian virus 40 DNA sequences in DNA of healthy adult mice derived from preimplantation blastocysts injected with viral DNA, Proc. Natl. Acad. Sci. USA, 71, 1250, 1974. 23. Anderson, W.F. et al. Replication and expression of thymidine kinase and human globin genes microinjected into mouse fibroblasts, Proc. Natl. Acad. Sci. USA, 77, 5399, 1980. 24. Capecchi, M.R. High efficiency transformation by direct microinjection of DNA into cultured mammalian cells, Cell, 22, (Pt 2), 479, 1980. 25. Gordon, J.W. et al. Genetic transformation of mouse embryos by microinjection of purified DNA, Proc. Natl. Acad. Sci. USA, 77, 7380, 1980. 26. Brinster, R.L. et al. Somatic expression of herpes thymidine kinase in mice following injection of a fusion gene into eggs, Cell, 27 (Pt 2), 223, 1981. 27. Janus, C., Chishti, M.A., and Westaway, D. Transgenic mouse models of Alzheimer’s disease, Biochim. Biophys. Acta, 1502, 63, 2000.

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28. Maguire-Zeiss, K.A. and Federoff, H.J. Convergent pathobiologic model of Parkinson’s disease, Ann. NY Acad. Sci., 991, 152, 2003. 29. Sathasivam, K. et al. Transgenic models of Huntington’s disease, Philos. Trans. R. Soc. Lond. B Biol. Sci., 354, 963, 1999. 30. Bates, G. Huntington aggregation and toxicity in Huntington’s disease, Lancet, 361, 1642, 2003. 31. Shibata, N. Transgenic mouse model for familial amyotrophic lateral sclerosis with superoxide dismutase-1 mutation, Neuropathology, 21, 82, 2001. 32. Jankowsky, J.L. et al. Transgenic mouse models of neurodegenerative disease: opportunities for therapeutic development, Curr. Neurol. Neurosci. Rep., 2, 457, 2002. 33. Wong, P.C. et al. Genetically engineered mouse models of neurodegenerative diseases, Nat. Neurosci., 5, 633, 2002. 34. Hsiao, K.K. et al. Age-related CNS disorder and early death in transgenic FVB/N mice overexpressing Alzheimer amyloid precursor proteins, Neuron, 15, 1203, 1995. 35. Evans, M.J. and Kaufman, M.H. Establishment in culture of pluripotential cells from mouse embryos, Nature, 292, 154, 1981. 36. Martin, G.R. Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells, Proc. Natl. Acad. Sci. USA, 78, 7634, 1981. 37. Robertson, E. et al. Germ-line transmission of genes introduced into cultured pluripotential cells by retroviral vector, Nature, 323, 445, 1986. 38. Isola, L.M. and Gordon, J.W. Transgenic animals: a new era in developmental biology and medicine, Biotechnology, 16, 3, 1991. 39. Balling, R. et al. Craniofacial abnormalities induced by ectopic expression of the homeobox gene Hox-1.1 in transgenic mice, Cell, 58, 337, 1989. 40. Beddington, R.S. et al. An in situ transgenic enzyme marker for the midgestation mouse embryo and the visualization of inner cell mass clones during early organogenesis, Development, 106, 37, 1989. 41. Palmiter, R.D. et al. Metallothionein–human GH fusion genes stimulate growth of mice, Science, 222, 809, 1983. 42. Iwamoto, T. et al. Aberrant melanogenesis and melanocytic tumour development in transgenic mice that carry a metallothionein/ret fusion gene, EMBO J., 10, 3167, 1991. 43. Mehtali, M., LeMeur, M., and Lathe, R. The methylation-free status of a housekeeping transgene is lost at high copy number, Gene, 91, 179, 1990. 44. Choi, T. et al. A generic intron increases gene expression in transgenic mice, Mol. Cell Biol., 11, 3070, 1991. 45. O’Gorman, S., Fox, D.T., and Wahl, G.M. Recombinase-mediated gene activation and site-specific integration in mammalian cells, Science, 251, 1351, 1991. 46. Lasko, P.F. Molecular movements in oocyte patterning and pole cell differentiation, Bioessays, 14, 507, 1992. 47. Kang, J. et al. The precursor of Alzheimer’s disease amyloid A4 protein resembles a cell-surface receptor, Nature, 325, 733, 1987. 48. De Strooper, B. and Annaert, W. Proteolytic processing and cell biological functions of the amyloid precursor protein, J. Cell Sci., 113 (Pt 11), 1857, 2000. 49. Tang, K. et al. Identification of a novel alternative splicing isoform of human amyloid precursor protein gene, APP639, Eur. J. Neurosci., 18, 102, 2003. 50. Golde, T.E. et al. Expression of beta amyloid protein precursor mRNAs: recognition of a novel alternatively spliced form and quantitation in Alzheimer’s disease using PCR, Neuron, 4, 253, 1990.

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51. Sandbrink, R., Masters, C.L., and Beyreuther, K. Beta A4-amyloid protein precursor mRNA isoforms without exon 15 are ubiquitously expressed in rat tissues including brain, but not in neurons, J. Biol. Chem., 269, 1510, 1994. 52. Clark, M.J. et al. MRC OX-2 antigen: a lymphoid/neuronal membrane glycoprotein with a structure like a single immunoglobulin light chain, EMBO J., 4, 113, 1985. 53. Rockenstein, E. et al. The neuroprotective effects of Cerebrolysin™ in a transgenic model of Alzheimer’s disease are associated with improved behavioral performance, J. Neural Trans., 110, 1313, 2003. 54. Quon, D. et al. Formation of beta-amyloid protein deposits in brains of transgenic mice, Nature, 352, 239, 1991. 55. Moran, P.M. et al. Age-related learning deficits in transgenic mice expressing the 751-amino acid isoform of human beta-amyloid precursor protein, Proc. Natl. Acad. Sci. USA, 92, 5341, 1995. 56. Higgins, L.S. et al. Early Alzheimer disease-like histopathology increases in frequency with age in mice transgenic for beta-APP751, Proc. Natl. Acad. Sci. USA, 92, 4402, 1995. 57. Games, D. et al. Alzheimer-type neuropathology in transgenic mice overexpressing V717F beta-amyloid precursor protein, Nature, 373, 523, 1995. 58. Irizarry, M.C. et al. Abeta deposition is associated with neuropil changes, but not with overt neuronal loss in the human amyloid precursor protein V717F (PDAPP) transgenic mouse, J. Neurosci., 17, 7053, 1997. 59. Irizarry, M.C. et al. APPSw transgenic mice develop age-related A beta deposits and neuropil abnormalities, but no neuronal loss in CA1, J. Neuropathol. Exp. Neurol., 56, 965, 1997. 60. Hsiao, K. et al. Correlative memory deficits, Abeta elevation, and amyloid plaques in transgenic mice, Science, 274, 99, 1996. 61. Westerman, M.A. et al. The relationship between Abeta and memory in the Tg2576 mouse model of Alzheimer’s disease, J. Neurosci., 22, 1858, 2002. 62. Andra, K. et al. Expression of APP in transgenic mice: a comparison of neuronspecific promoters, Neurobiol. Aging, 17, 183, 1996. 63. Chishti, M.A. et al. Early-onset amyloid deposition and cognitive deficits in transgenic mice expressing a double mutant form of amyloid precursor protein 695, J. Biol. Chem., 276, 21562, 2001. 64. Kulnane, L.S. and Lamb, B.T. Neuropathological characterization of mutant amyloid precursor protein yeast artificial chromosome transgenic mice, Neurobiol. Dis., 8, 982, 2001. 65. Aguzzi, A. et al. Transgenic and knock-out mice: models of neurological disease, Brain Pathol., 4, 3, 1994. 66. Guo, Q. et al. Increased vulnerability of hippocampal neurons to excitotoxic necrosis in presenilin-1 mutant knock-in mice, Nat. Med., 5, 101, 1999. 67. Siman, R. et al. Presenilin-1 P264L knock-in mutation: differential effects on abeta production, amyloid deposition, and neuronal vulnerability, J. Neurosci., 20, 8717, 2000. 68. Hayashi, H. et al. Cholesterol is increased in the exofacial leaflet of synaptic plasma membranes of human apolipoprotein E4 knock-in mice, Neuroreport, 13, 383, 2002. 69. Zheng, H. et al. Beta-amyloid precursor protein-deficient mice show reactive gliosis and decreased locomotor activity, Cell, 81, 525, 1995. 70. Mayford, M. et al. Control of memory formation through regulated expression of a CaMKII transgene, Science, 274, 1678, 1996.

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71. Oddo, S. et al. Triple-transgenic model of Alzheimer’s disease with plaques and tangles: intracellular Abeta and synaptic dysfunction, Neuron, 39, 409, 2003. 72. Oddo, S. et al. Amyloid deposition precedes tangle formation in a triple transgenic model of Alzheimer’s disease, Neurobiol. Aging, 24, 1063, 2003. 73. Andorfer, C. et al. Hyperphosphorylation and aggregation of tau in mice expressing normal human tau isoforms, J. Neurochem., 86, 582, 2003. 74. Carlson, G.A. et al. Genetic modification of the phenotypes produced by amyloid precursor protein overexpression in transgenic mice, Hum. Mol. Genet., 6, 1951, 1997. 75. Brinster, R.L. et al. Factors affecting the efficiency of introducing foreign DNA into mice by microinjecting eggs, Proc. Natl. Acad. Sci. USA, 82, 4438, 1985. 76. Taketo, M. et al. FVB/N: an inbred mouse strain preferable for transgenic analyses, Proc. Natl. Acad. Sci. USA, 88, 2065, 1991. 77. Costantini, F. and Lacy, E. Introduction of a rabbit beta-globin gene into the mouse germ line, Nature, 294, 92, 1981. 78. Taylor, L.D. et al. A transgenic mouse that expresses a diversity of human sequence heavy and light chain immunoglobulins, Nucleic Acids Res., 20, 6287, 1992. 79. Strouboulis, J., Dillon, N., and Grosveld, F. Developmental regulation of a complete 70-kb human beta-globin locus in transgenic mice, Genes Dev., 6, 1857, 1992. 80. Pieper, F.R. et al. Efficient generation of functional transgenes by homologous recombination in murine zygotes, Nucleic Acids Res., 20, 1259, 1992. 81. Choi, T.K. et al. Transgenic mice containing a human heavy chain immunoglobulin gene fragment cloned in a yeast artificial chromosome, Nat. Genet., 4, 117, 1993. 82. Brinster, R.L. et al. Introns increase transcriptional efficiency in transgenic mice, Proc. Natl. Acad. Sci. USA, 85, 836, 1988. 83. Palmiter, R.D. et al. Heterologous introns can enhance expression of transgenes in mice, Proc. Natl. Acad. Sci. USA, 88, 478, 1991. 84. Chada, K. et al. Specific expression of a foreign beta-globin gene in erythroid cells of transgenic mice, Nature, 314, 377, 1985. 85. Krumlauf, R., Hammer, R.E., Tilghman, S.M., and Brinster, R.L. Developmental regulation of alpha-fetoprotein genes in transgenic mice, Mol. Cell Biol., 5, 1639, 1985. 86. Townes, T.M. et al. Erythroid-specific expression of human beta-globin genes in transgenic mice, EMBO J., 4, 1715, 1985. 87. Hammer, R.E. et al. Diversity of alpha-fetoprotein gene expression in mice is generated by a combination of separate enhancer elements, Science, 235, 53, 1987. 88. Behringer, R.R. et al. Synthesis of functional human hemoglobin in transgenic mice, Science, 245, 971, 1989.

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14

Generation of Amyloid Precursor Protein Knockout Mice Hui Zheng

CONTENTS 14.1 Introduction 14.2 Experimental Procedures 14.2.1 Construction of APP Gene Targeting Vector 14.2.2 Embryonic Stem Cells: Culturing and Maintenance 14.2.2.1 Growing SNL Cells and Preparation of Feeders 14.2.2.2 Culturing and Electroporating Embryonic Stem Cells 14.2.3 Selection of Homologous Recombinant Clones 14.2.3.1 Selection of Recombinant Colonies 14.2.3.2 Expansion and Duplication of Recombinant Clones 14.2.3.3 Freezing ES Cells in 96-Well Plates 14.2.3.4 Mini-Southern Analysis of ES Clones to Identify Gene Targeting Events 14.2.3.5 Preparation of Gene-Targeted Clones for Blastocyst Injection 14.2.4 Generation of Gene (APP) Knockout Mice 14.2.4.1 Microinjection, Assessment of Chimerism, and Test for Germline Transmission 14.2.4.2 Breeding and Generation of Homozygous APP Knockout Mice Acknowledgments References

14.1 INTRODUCTION Alzheimer’s disease (AD) is the most common cause of dementia in the aged population. It is characterized pathologically by the deposition of β-amyloid plaques, the accumulation of neurofibrillary tangles and the losses of neurons and synapses in selected areas of the brain. Senile plaques are extracellular deposits of heterogeneous substances of which the major components are the 40 to 42 amino acid 0-8493-2245-6/05/$0.00+$1.50 © 2005 by CRC Press

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peptides referred to as β-amyloid peptides (Aβ) derived by proteolytic cleavage of the amyloid precursor protein (APP). Approximately 10% of AD cases are familial and cosegregate with autosomal dominant inheritance of mutations in genes including APP,1 presenilin 1 (PS1),2 and the PS1 homolog, presenilin 2 (PS2).3 APP is an integral membrane glycoprotein consisting of an extracellular domain, a single transmembrane domain and a short cytoplasmic tail. About two-thirds of the Aβ peptide is extracellular and the remaining sequence is embedded in the membrane. The APP gene located on the long arm of chromosome 21 spans approximately 400 kb and contains at least 18 exons.4,5 Alternative splicing generates APP mRNAs encoding several isoforms that range from 365 to 770 amino acid residues. The major Aβ peptide-encoding proteins are 695, 751, and 770 amino acids (referred to as APP695, APP751, and APP770). APP751 and APP770 contain domains homologous to the Kunitz-type serine protease inhibitors (KPIs) and are expressed in most tissues examined. The APP695 isoform lacks the KPI domain and is predominantly expressed in neurons. APP is processed by at least three proteinases termed α-, β-, and γ-secretases. α-Secretase cleavage occurs inside the Aβ domain, thus precluding intact Aβ peptide formation. The identity of the α-secretase has not been unambiguously established but the ADAM family of metalloproteases known as ADAM 10 and 17 (the latter is also known as TACE) has been reported to exhibit the activities.6,7 β-Secretase (BACE1) cleaves APP at the amino terminus of Aβ. The enzyme has been cloned and its characteristics extensively studied.8 These processing events generate large soluble APP derivatives (called APPsα and APPsβ, respectively). Following the extracellular cleavages, γ-secretase processes APP at the carboxyl-terminus of Aβ to produce either a 3-kDa product (p3 in combination with the α-secretase) or Aβ (in concert with BACE1 cleavage), respectively. The γ-secretase activity seems to be executed by a high molecular weight complex of which presenilin (PS1 or PS2) is an essential component.9 In addition to the γ-cleavage that yields Aβ40 and Aβ42, presenilin-dependent proteolysis also occurs at the Leu 49–Val 50 position (counted from Aβ) downstream of the γ-site and near the membrane intracellular boundary (termed ε-site). This ε-cleavage releases an APP intracellular domain (AICD).9,10 Because of the central role of APP in AD pathogenesis, the expression patterns, biochemical properties, and potential functions of APP have been the subjects of intensive studies in vitro. Within the nervous system, APP is present on cell surfaces and in axons, dendrites, and vesicles.11 It undergoes rapid anterograde transport12–14 and is targeted to the synaptic sites of both central and peripheral nervous systems.15–17 The major activities reported include vesicular transport, transcriptional regulation, synaptogenesis, trophic activity, and cell motility.9 The extracellular domain of APP has been proposed to exhibit neurotrophic, synaptogenic, and growth-promoting properties.18–21 Two distinct activities of the APP intracellular domain (AICD) have been reported. First, it binds to kinesin molecular motors and has been shown to regulate axonal transport of prepackaged vesicles.22–24 Second, the APP IC interacts with various adaptor proteins including Fe65, X11, and mDAB1.9 Following presenilin-dependent intramembrane proteolysis, AICD can be released, and the AICD/Fe65 complex can translocate to the

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nucleus25,26 where, in concert with the histone acetyltransferase Tip60, it functions as a potent transcriptional regulator.27 This processing and signaling mechanism is analogous to the Notch signaling pathway28 and several potential downstream targets have been proposed.29–31 Several disease-causing missense mutations in the human APP gene have been identified.32 Among these, the KM-to-NL double (Swedish or APPsw) mutation is present immediately upstream of the BACE cleavage site and seems to enhance the BACE1 cleavage to increase both Aβ40 and Aβ42 levels. The Val mutation downstream of the γ-secretase cleavage site results in a specific increase of the Aβ42 peptide.33,34 These mutations apparently affect the γ-secretase activity that absolutely requires PS1 and PS2.35–37 Consistent with this activity, mutations in presenilins lead to a specific increase in Aβ42 production,38 although the molecular mechanism of this apparent “gain-of-misfunction” effect is not well understood. Interestingly, the PS1L166P familial AD mutation results in an increase in Aβ42 (γ-cleavage) and a decrease in AICD (ε-cleavage).10 Likewise, the same mutation leads to impaired Notch intracellular domain (NICD) production.10 The effect of the presenilin AD mutation may be partial loss of function with respect to the signaling pathways of Notch and, potentially, APP. To understand the in vivo function of APP and its processing products, we generated an APP null mutation in mice using gene targeting in embryonic stem (ES) cells.39 This technology was developed based on (1) the successful isolation of permanent in vitro ES cell lines from preimplantation blastocyst stage embryos40; (2) documentation indicating that these cell lines, after sustained in vitro manipulation and growth, were capable of recolonizing embryos and contributing to the germlines41; and (3) a demonstration that many mammalian cells42 including ES43 have a remarkable ability that allows transfected foreign DNA to locate and recombine with its homologous chromosomal counterpart.42,43 This process, referred to as homologous recombination or gene targeting, allows virtually any genetic modification of endogenous genes such as inactivation of a gene of interest (gene knock-out), subtle and precise genetic modification (gene knock-in) and large chromosomal deletions (chromosome engineering).44 This chapter discusses only procedures and considerations related to the generation of the APP knockout mice. Readers interested in general aspects of gene targeting technology are referred to reviews in References 45 through 48.

14.2 EXPERIMENTAL PROCEDURES The generation of APP knockout mice requires the following steps: 1. Construct an APP gene targeting vector that, upon homologous recombination into the endogenous APP locus, can completely inactivate the gene. 2. Introduce the vector into ES cells and identify ES clones with the disrupted APP gene. 3. Microinject the targeted APP clones into mouse blastocyst stage embryos, transplant the injected embryos into a foster mother, and bring them to

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term to generate chimeric mice with contributions from both donor ES cells and host embryos. 4. Identify and breed germline chimeras capable of transmitting the APPtargeted allele to their offspring. The offspring will contain one copy of the wild-type allele and one copy of the APP-targeted allele and are called APP heterozygous (APP+/–) mice. 5. Interbreed the APP heterozygous mice to create animals that are either wild-type (APP+/+), APP+/–, or homozygous for the APP mutant allele (APP–/–). These procedures are illustrated in Figure 14.1 and described below.

14.2.1 CONSTRUCTION

OF

APP GENE TARGETING VECTOR

A typical gene targeting vector of replacement type consists of two segments of sequences homologous to the endogenous gene. The genomic sequences between the two segments that encode essential functional units of the gene are deleted and replaced by a selectable marker. The most common marker is the neomycin resistance gene (neo). Selection of transfected cells with geneticin antibiotic (G418) allows identification of total integration events, including both homologous recombination and random integration. In most circumstances, random integration is the predominant event. Many factors can influence the ratio of homologous recombination to random integration, for example, target locus, length of homology, and degree of polymorphism between the vector sequence and the endogenous chromosomal DNA. As a general rule, given a target locus, the longer the homologous sequence, the higher the targeting frequency. To minimize polymorphic variations and optimize homologous recombination, the DNA in the targeting vector should be isolated from the same strain as that of the ES cells. In addition to positive selection using the neomycin resistance gene, a negative selectable gene, HSV-tk, can be added at the end of the linearized targeting vector. Incorporation of HSV-tk by random integration, not homologous recombination (lost due to homologous crossover, Figure 14.2a), renders the cells sensitive to antiviral agents 1-(2′-deoxy, 2′-fluoro-β-δ-arabinofuranosyl)-5-iodouracil (FIAU) and gancyclovir and, as a result, enriches gene-targeting events.49 Inactivation of the 400-kb APP gene which undergoes alternative splicing requires a careful design of the targeting vector to ensure that an APP null allele is created. We chose to delete the APP promoter and first exon on the assumption that the mutation would prevent APP gene transcription. If an alternative promoter and ATG initiation codon were used, transport of the protein to the membrane should have been impaired because the signal peptide would have been deleted. Using this rationale and the general gene-targeting principle described above, we used a 1.0-kb HindIII–PvuII fragment of the mouse APP promoter50 as a probe, and screened a genomic-phage library made from the 129Sv strain (the strain AB2.1 ES cells are derived from) of DNA. We isolated the APP promoter and surrounding regions

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1)

Electroporation

2)

Selection

3) Blastocyst injection

4) Chimera

Targeted progeny

5)

Wild-type progeny

APP+/-X APP+/-

APP+/+APP+/-APP/-

FIGURE 14.1 Gene targeting outline. (1) A gene-targeting construct (double helix) containing a genetic modification and a positive selectable marker (asterisk) is introduced into mouse embryonic stem (ES) cells by electroporation. (2) Cells that have integrated the genetargeting construct are resistant to drug selection due to the presence of the positive selectable marker in the genome. These cells form colonies (circles). Clones that result from accurate recombination at the target locus are recognized by Southern hybridization or PCR. (3) Targeted clones are injected into developing embryos at the blastocyst stage and embryos are implanted into the uteri of surrogate mothers. Mice that have contributions from the genetically modified ES cells are recognized by their chimeric coat colors because the blastocysts are derived from black mice (light circles), whereas the ES cells used for targeting are derived from a pigmented agouti mouse (dark circles). (4) Mouse strains are established by mating chimeras to black C57BL/6 mice. Germline transmission results in offspring that are agouti color consisting of one half wild-type mice and the other half targeted, heterozygous mice. (5) Heterozygous APP+/– mice are interbred to produce mice that are wild-type (APP+/+), heterozygous (APP+/–), or homozygous (APP–/–) for the targeted APP allele. (Reprinted from Mills, A.A. and Bradley, A. Trends Genet., 17, 331, 2001. With permission from Elsevier.)

and constructed an APP targeting vector by ligating a 1.4-kb BglII–XhoI fragment from the 5′ portion of the APP promoter and a 7.1-kb XhoI–BglII fragment from the 3′ portion of exon 1. A 1.5-kb XhoI–SalI PGK–neo cassette was inserted between the segments, deleting a 3.8-kb XhoI–XhoI fragment encoding the APP promoter and the first exon. The PGK–neo cassette was inserted in an opposite orientation to avoid potential transcriptional read-through to the APP sequence by the PGK promoter.

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a. R Targeting Vector HSV-tk

PGKneo R

X

ATG

B Pr

9.0 kb

B N

R

R Wild-type APP Locus

E1

9.5 kb

Homologous Recombination R

X

B

R

B N

R Targeted APP Locus

PGKneo

6.5 kb

b.

2.1

AB

76

9.0 kb

3

12

4

17

6

19

kb

9.0

6.5

FIGURE 14.2 Targeted disruption of the APP gene in ES cells. (a) The targeting vector contains the following, from left to right: a 1.4-kb segment preceding the APP promoter (hatched 5′ rectangle); a PGK–neo expression cassette inserted in an orientation opposite that of the APP gene; a fragment of 7.1 kb homologous to the first intron of the APP gene (hatched 3′ rectangle); and a HSV-tk gene (closed box) for negative selection with FIAU. Probes used (black boxes below physical map) for detecting targeted events are 1.0-kb XbaI-BglII (5′) and 0.8-kb BglII-NcoI (3′) probes. Digestion with EcoRI is used to separate the wild-type and targeted APP alleles. R = EcoRI. X = XbaI. B = BglII. N = NcoI. Pr = mouse APP promoter. E1 = exon 1 of the mouse APP gene. (b) Southern blot analysis of representative targeted clones (76, 123, 174, 196) using the 5′-probe. AB2.1 wild-type ES cells.

The final vector was made by ligating a 2-kb XhoI fragment of pMC–TK to the end of the 7.1-kb homologous sequence (Figure 14.2a). The targeting vector was linearized by digestion with NotI prior to electroporating into ES cells. In addition to the DNA sequences used for vector construction, at least one and preferably two DNA fragments of 300 to 1000 bp outside the vector should be isolated and tested against a genomic blot to make sure that they are free of repetitive sequences. They will be used as probes to screen for gene-targeting events from total recombinant clones by Southern blot analysis. Because the probe does not hybridize to the vector DNA, only the endogenous allele, either wild-type or disrupted by the vector, will be recognized on a Southern blot. To properly identify the gene-targeted clones, a restriction digestion scheme should be in place to distinguish the wild-type and targeted alleles upon hybridization with the probe. For targeting the APP locus, we used EcoRI digestion because introduction of a new EcoRI site by PGK–neo insertion alters the restriction patterns of the targeted allele (Figure 14.2).

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Hybridization with the 5′-probe creates a 9.0-kb wild-type band and a 6.5-kb targeted band that can be separated easily and distinguished by Southern blotting. The homologous recombination event can be further confirmed using the 3′-probe that yields 9.5- and 9.0-kb fragments by the wild-type and targeted alleles, respectively (not shown).

14.2.2 EMBRYONIC STEM CELLS: CULTURING

AND

MAINTENANCE

The key requirement to successful generation of gene knockout mice is that both the parental and targeted ES clones retain their totipotencies to contribute to the somatic lineage and germline following microinjection into host embryos. Progressive culturing and cloning of ES cells — necessary procedures for gene targeting experiments — may produce ES cells that have lost their totipotency but are morphologically indistinguishable from normal ES cells. These cells, however, can only produce low-percentage chimeras that are not capable of transmitting through the germline. Therefore, maintaining ES cells under optimal conditions is extremely important. The culture of ES cells requires a complex mixture of growth factors provided by fetal bovine serum and mitotically inactivated fibroblast feeder cells. The right combination of ES and feeder pair is essential to ensure germline transmission. The AB1 and AB2.1 ES cells growing on clonal SNL76/7 feeder cell layers represent an excellent system (reviewed in Reference 51). Both ES lines are derived from the 129Sv strain of blastocysts and the feeder cells are generated by transfecting the STO fibroblasts with a neomycin marker and leukemia inhibitory factor (LIF, an essential component for keeping ES cells in an undifferentiated state), hence the name SNL. The detailed culture procedure is described below. 14.2.2.1 Growing SNL Cells and Preparation of Feeders All tissue culture reagents can be purchased from Gibco/Invitrogen (Carlsbad, CA) unless otherwise noted. Disposable plastic ware should be used for culturing and processing the cells. Although products from various suppliers can be used, we use Costar plates (Corning, Inc., NY). SNL medium — 91% dimethyl sulfoxide (DMSO, high glucose, no pyruvate), 7% fetal bovine serum (FBS), 1% L-glutamine, 1% antibiotics (penicillin and streptomycin mix). SNL cells — These are easy to culture and fast growing. They can be thawed and cultured under routine tissue culture conditions using the above SNL medium. Split the cells every 3 days at a ratio of 1:8. Feeder preparation — Add 260 µl of 0.5 mg/ml mitomycin C solution* (Sigma, St. Louis, MO) to a 100-mm dish containing confluent SNL cells in 12 ml medium. Incubate the cells at 37ºC for 3 hr. Coat appropriate dishes where the feeder cells are to be plated with a 1% gelatin solution. Remove gelatin before plating the feeder layer. Wash the mitomycin C-treated cells twice with PBS. Trypsinize the cells using 2.5% trypsin/EDTA at 37oC for 5 min. Spin down and resuspend the cells in SNL * Add 4 ml PBS to a 2-mg bottle. Block the solution from light. Use within 2 weeks.

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medium. Count the cells and dilute to 3.5 × 105/ml. Plate 12 ml on a 100-mm gelatintreated dish, 4 ml/60-mm dish, 2 ml/6-well dish, and 0.5 ml/24-well dish. For 96-well plates, dilute the cells to 1.15 × 105/ml and plate 150 µl/well using a multichannel pipettor. Feeder Maintenance — The feeder cells can be maintained in a 37ºC incubator for at least 5 weeks. Feed the cells with fresh SNL medium once every 2 weeks after plating. In general, older feeder plates are better than new ones and 3- to 4-week-old plates are best. For one gene targeting experiment, 5 × 100 mm and 10 × 96-well plates are needed and should be prepared prior to electroporation. Feeder plates of other sizes are used for expanding ES stocks and selected clones and should be prepared as planned. 14.2.2.2 Culturing and Electroporating Embryonic Stem Cells ES cell medium — 83% DMSO (high glucose, no pyruvate), 15% FBS, 1% L-glutamine, 1% antibiotics (penicillin and streptomycin mix), 1% β-mercaptoethanol. Serum testing — Use of high-quality FBS is crucial for optimal culturing of ES cells. The serum should be tested and compared against a control with a successful track record. Sera from several companies and lots should be tested. We have good success with HyClone products (Logan, UT) in general. For serum testing, plate ES cells onto 60-mm feeder plates at 2000 cells/plate. Five plates should be used for each batch of serum: four with 15% serum and one with 30%. Incubate plates at 37ºC for 10 to 14 days. Use one 15% plate each for daily examination of colony size and morphology. At the end of incubation, the remaining plates should be washed twice with PBS and stained with methylene blue solution (2% w/v in 70% EtOH) for 5 min. The remaining plates will be washed once with 70% EtOH and allowed to dry. The number of colonies on each plate will be counted. Average the three 15% serum counts for each batch. Compare the 30% and 10% values of each serum lot with those of the control serum. Select the batch that yields a plating efficiency equal to or greater than the appropriate control sample and shows no toxicity at 30%. A large stock should be purchased to avoid frequent testing. Alternatively, ESqualified sera are available from several companies. The ones specifically tested for the AB1/AB2.1-SNL system can be used to culture the ES cells. ES cell culturing — The derivation, general morphology and growth properties of ES cells are illustrated in Chapter 4 of Robertson’s book.52 The doubling time of ES cells is in the range of 18 to 24 hr. The following rules should be followed when culturing AB1 or AB2.1 ES cells: 1. The cells should grow on a feeder layer with serum-tested ES medium at relatively high frequency (30 to 80% confluent). They should be split every 3 to 5 days. Always feed the cells with fresh medium 2 to 3 hr prior to passaging. 2. The cells have a high tendency to secrete acidic material and color the medium yellow. Extremely acidic pH is harmful to the cells and therefore the medium should be replaced every 2 days under normal growth conditions or whenever the color becomes yellowish. All medium used should

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be orange (pH 7.4). Old, pinkish medium has a pH of 7.6 or higher and should be avoided. 3. The ES cells grow in patches. To maintain the cells in an undifferentiated state, it is extremely important to adequately break them with trypsin. Use 2.5% trypsin–EDTA, incubate the cells at 37ºC for 15 min, and pipette the cells vigorously five to eight times after neutralizing the trypsin with an equal volume of ES medium. 4. Long-term passaging of ES cells will reduce their germline transmission potential. A large stock of ES cells at relatively early passages (below 20) should be frozen in liquid nitrogen. A freshly thawed vial should be used for each gene targeting experiment. Freezing and thawing ES cells — The general rule of slow freezing and quick thawing applies. ES cells to be frozen should be grown in log phase and fed 2 to 3 hr before trypsinization. Wash the plates twice with PBS. Add 2.5% trypsin–EDTA (1 ml/60-mm dish). Incubate at 37ºC for 15 min. Add an equal volume of ES medium. Pipette cells vigorously five to eight times with a Pasteur pipette. Transfer the solution to a 15-ml conical tube and spin the cells down using a low speed centrifuge at 1000 rpm for 7 min. Resuspend cells in ES medium at 1 to 2 × 107/ml. Add dropwise an equal volume of 2X freezing medium (50% DMEM, 30% FBS, 20% DMSO) while mixing. Aliquot into 1-ml cryotubes. Place the tubes in a polystyrene box and cool in a –80ºC freezer overnight. Transfer the vials to liquid nitrogen the next day. Thawing — Retrieve a frozen vial from storage and place directly in a 37ºC water bath until the solution is completely thawed. Sterilize the vial with 70% EtOH and transfer the cells to a 15-ml conical tube containing prewarmed ES medium. Spin cells down; resuspend in 4 ml ES medium and plate onto a 60-mm feeder dish. Electroporation — The targeting vector (25 µg) should be linearized at one end by restriction enzyme digestion and resuspended in Tris-EDTA, pH 7.5, at 1 µg/µl. Low-passage (<20) ES cells should be grown in log phase and fed 2 to 3 hr prior to electroporation. The cells are trypsinized and resuspended in PBS at 1.2 × 107/ml using the protocol described above. Add DNA to 800 µl of cells and mix by inverting. Transfer the mixture to a Gene Pulser cuvette with a 4-mm electrode gap (Bio-Rad Laboratories, Hercules, CA) and electroporate at 230 V, 500 µF using a Bio-Rad Gene Pulser apparatus with a capacitance extender. The pulse time under this condition is 0.6 to 0.8 msec. Transfer the treated cells to a bottle containing 60 ml ES medium. Evenly distribute onto 5 × 100 mm feeder plates at 12 ml/plate.

14.2.3 SELECTION

OF

HOMOLOGOUS RECOMBINANT CLONES

14.2.3.1 Selection of Recombinant Colonies Twenty-four hours after electroporation, G418 (200 µg/ml active ingredient, from 1000X stock) is added to the plates. If HSV-tk is present in the vector, FIAU is added at a final concentration of 0.4 µM. Change media every other day for 8 to

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10 days. Nontransfected cells start to die at 4 to 5 days with the appearance of G418or G418/FIAU-resistant colonies soon afterward. Under the electroporation conditions described above, we expect the transfection efficiency to be in the range of 1 × 10–4 to 5 × 10–3. Therefore, with 1 × 107 cells electroporated, approximately 1000 to 5000 total G418-resistant colonies should be present. With FIAU selection, we usually achieve a two- to fivefold enrichment and thus the number of G418/FIAU double-resistant clones is expected to be reduced by two- to fivefold. Most of the colonies should have round shapes and be densely packed with darker central cores. They should be large enough to be picked 8 to 10 days after G418 selection. Prolonged growth of ES colonies leads to differentiation and should be avoided. Differentiated cells can be distinguished from normal ES cells since they are usually loosely packed and exhibit different morphologies. 14.2.3.2 Expansion and Duplication of Recombinant Clones The G418- or G418/FIAU-resistant colonies are picked for further expansion and mini-Southern analysis.53 Place 25 µl of 0.25% trypsin–EDTA into each well of a round-bottom 96-well plate using a multichannel pipette. Wash plates with colonies twice with PBS. Leave 1 to 2 ml in plates the second time. Using a 200-µl pipette set at 20 µl, pick colonies from the washed plates and transfer to each trypsincontaining well. Carefully mark the wells to make sure each well contains only one colony. Our picking speed is ~100 to 150 colonies/hr. Therefore, when a 96-well is completed, enough time has elapsed to allow complete trypsinization and no further incubation is necessary. Multichannel pipettes will be used for most of the procedures from this point on. One row at a time from earlier-picked colonies, add 25 µl of ES medium per well and pipette up and down a dozen times to disaggregate the cells. Transfer the entire solution to a 96-well feeder plate. Repeat with another dish. We normally pick 400 colonies per experiment to ensure positive identification of a minimum of two to four targeted clones (assuming a targeting frequency of 1/100). Change to fresh ES media containing G418 the next day and every other day afterward and allow colonies to grow (~4 to 6 days). ES cells should be visible under a light microscope on the third day. The average rate of colony recovery (percentage of wells with medium color change) is around 70 to 80%. The ES clones will then be duplicated: one set to gelatin plates for DNA isolation and mini-Southern analysis (see below) and the other set to feeder plates for freezing and storage. Feed the wells 2 to 3 hours before treatment. Wash cells twice with PBS. Add 50 µl 0.25% trypsin–EDTA per well and incubate at 37ºC for 10 min. Add 100 µl ES medium to each well and mix by pipetting up and down about 10 times. Evenly transfer the solution to one gelatin-coated plate and one feeder plate and fill each well with 150 µl additional ES medium. 14.2.3.3 Freezing ES Cells in 96-Well Plates The feeder plate will be frozen as the master. Trypsinize the plate as above. Neutralize and disaggregate the cells with 50 µl ES medium. Add dropwise 100 µl 2X freezing

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medium (50% DMEM, 30% FBS, 20% DMSO) to each well, followed by 50 µl sterilized (0.2 µm filtered) light paraffin oil to prevent degassing and evaporation. Tape the lid to the bottom of the plate and place in a polystyrene box. Cover with lid, freeze, and store at –80ºC. 14.2.3.4 Mini-Southern Analysis of ES Clones to Identify Gene Targeting Events Allow cells on gelatin plates continue to grow until most wells are near confluency. Wash twice with PBS. Add 50 µl of lysis buffer per well (10 mM Tris, pH 7.5, 10 mM EDTA, 10 mM NaCl, 0.5% sarcosyl, and 1 mg/ml proteinase K added fresh). Incubate overnight at 60ºC in a humid atmosphere (e.g., a sealed container with a presoaked sponge). The next day, prepare a mix of NaCl and EtOH (1.5 µl of 5 M NaCl to 100 µl of cold absolute ethanol; salt will precipitate, so mix well). Add 100 µl per well. Let the plate stand at room temperature 30 min. Gently invert the plate to discard the solution. Wash twice with 70% ethanol. After the final washing, invert the plate and let it semi-dry for 15 min. While the plate dries, prepare a restriction digest cocktail containing 1X restriction buffer, 1 mM spermidine, 100 µg/ml bovine serum albumin and 10 to 15 units of enzyme per well). Add 30 µl of restriction digest cocktail to each well and incubate at 37ºC overnight in a humid atmosphere. Add 4 to 5 µl of loading dye per well the next day and load onto a large 0.7 to 1.0% agarose gel containing three rows (33 wells per row). Each gel can hold 96 samples (one 96-well) plus one molecular weight marker per row. Run gel at 80 V for 3 to 5 hr, depending on the predicted size differences. Proceed with standard Southern blot analysis. 14.2.3.5 Preparation of Gene-Targeted Clones for Blastocyst Injection The targeted clones identified by above mini-Southern analysis will be expanded for blastocyst injection. Feeder plates of 6 and 24 wells should be prepared. To retrieve cells from frozen 96-well plates, place plates directly from –80ºC freezer into 37ºC incubator. It takes 10 to 15 min for the wells in the center of the plate to thaw. After all the wells have thawed, carefully identify and mark the wells with targeted clones and remove the entire solution to a 24-well feeder plate pre-equilibrated with 2 ml of ES medium. It is important for maximum recovery to vigorously pipette the thawed cells and rinse once to dislodge them from the bottom of the plate where they settle during the freezing process. Change with fresh medium the second day. Split the cells after 4 or 5 days to 6-well feeder plates. If the cell density is low, split to two 24-wells. It is not advisable to grow ES cells too long without passaging. Cells grown on 6-well plates can be divided for several purposes: further splitting and expansion, freezing to make a permanent stock, and isolating DNA to doublecheck the genotype. Clones with good growth characteristics and morphology will

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be chosen for microinjection. The viability of cells after trypsinization determines the quality of chimeras. Badly maintained cultures make poor chimeras. The cloning efficiency at the time of microinjection is much more critical than for routine maintenance of the culture. The cells should be in log phase growth to ensure maximum viability post-trypsinization. This is best achieved by plating the cells at a high density 24 to 32 hr prior to injection. Cells from a 24-well plate are more than sufficient for a day of injection. Feed the cells 2 to 3 hr before splitting. After trypsinization for 15 min, add an equal volume of ES medium with 20 mM Hepes (pH 7.4). Add an equal volume of FBS and pipette vigorously five to eight times with a Pasteur pipette. Transfer to 1-ml screw cap tube and place on ice.

14.2.4 GENERATION

OF

GENE (APP) KNOCKOUT MICE

14.2.4.1 Microinjection, Assessment of Chimerism, and Test for Germline Transmission Microinjection of targeted ES cells to create chimeric mice is a complicated process and requires sophisticated instruments. It is usually done by a core facility and is not recommended for inexperienced researchers. This chapter will not discuss the procedure. Interested individuals are referred to Bradley’s article about production and analysis of chimeric mice (Chapter 5 in Robertson’s book).52 Pigmentation markers are commonly used to identify chimeras and to quantify the extent of the chimerism. A common strain of host embryo is C57BL/6 (black, aa). The ES cells are 129Sv (agouti coat color, AA). Chimerisms are seen as areas of brown hairs on black backgrounds in mice 10 days and older. The percentage of chimerism is a good prediction for germline transmission. A chimera with good chance of germline transmission should have a chimerism of 70% or higher. However, while a poor chimera most likely will not transmit through the germline, a high percentage chimera does not guarantee germline transmission. Usually high percentage male chimeras are used to test for germline transmission. There are two reasons: 1. Because the ES cells are derived from XY males, if the host embryo is a female, it is possible for the ES cells to convert the host from female to male. These sex-converted males are definitive germline transmitters if they are fertile. 2. In contrast to females that need to produce offspring one litter at a time, a male can simultaneously breed with multiple partners and a large number of offspring can be generated in a short time. This is particularly important if the chimera is a low germline transmitter, i.e., only a small percentage of pups are ES-derived. Coat color is again used as a convenient and convincing test for germline transmission. When the 129Sv (AA) and C57BL/6 (aa) mosaic chimeras are bred with C57BL/6 (aa) counterparts, the offspring can be either Aa (if 129Sv allele

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contributes to germline) or aa (if germ cells carry two C57BL/6 alleles) with regard to the pigmentation locus. Since agouti (A) is dominant over black (a), an Aa combination will confer agouti coat color and, therefore, germline transmission can be determined by the presence of agouti pups in the offspring. The resulting mice are on a mixed C57BL/6 and 129Sv background. Due to the convenience and fast breeding scheme, this is the most commonly used genetic background for initial phenotypic characterization of the mutant mice. 14.2.4.2 Breeding and Generation of Homozygous APP Knockout Mice Because only one ES allele is targeted, 50% of the agouti offspring are expected to carry the targeted allele and the other 50% wild-type. They can be distinguished, upon tail biopsy at weaning age, by Southern blot analysis described above (Figure 14.3a). However, it is advisable to develop a much faster and easier PCR method. This can facilitate the genotyping efficiency tremendously. In the case of APP, we designed the following trimer mix (Figure 14.3b):

a.

+/+

+/+

+/-

+/-

-/-

-/-

kb

9.0

6.5

b.

-/-

+/+

+/-

+/-

-/-

bp

470

250

FIGURE 14.3 Southern blot (a) and PCR (b) analysis of representative offspring from heterozygous mating of wild type (+/+), heterozygous (+/–), and homozygous (–/–) APP knockout mice.

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Primer 1 — APP wild-type sense oligo (2.2 kb into intron 1, deleted in targeted allele): 5′-CTG CTG CAG GTG GCT CTG CA-3′ Primer 2 — APP wild-type/targeted antisense oligo (2.45kb into intron 1, present in both alleles): 5′-CAG CTC TAT ACA AGC AAA CAA G-3′ Primer 3 — Neo antisense oligo (in proximal neo promoter 2540–2522, specific to targeted allele): 5′-CCA TTG CTC AGC GGT GCT GTC CAT-3′ Expected band pattern — APP wild-type allele (primers 1 and 2): 250 bp; APP targeted allele (primers 2 and 3): 470 bp; heterozygous: 250 bp + 470 bp Heterozygous mice identified can be crossbred to yield mice that are wild-type (APP+/+), heterozygous (APP+/–), or homozygous (APP–/–) for the targeted APP allele (Figure 14.3). If the homozygous mice are viable as in the case of APP, the expected ratios are 25, 50, and 25%, respectively. If embryonic or early postnatal lethality arises as is the case for PS1 knockout,54 homozygous mice may not be recovered at weaning age. Genotyping and characterization at an earlier age or during embryonic development are required. After the homozygous knockout mice are identified, it is important to perform expression analysis to determine whether a null allele has been created. This can be done by various methods — the most convincing are Northern and Western blotting (Figure 14.4). By deleting the promoter and the first exon of APP, we were able to eliminate the APP transcript (Figure 14.4a) and protein (Figure 14.4b) completely. The homozygous APP-null mice and their littermate controls are subjected to various biochemical, immunohistochemical, and electrophysiological studies.39,55–57 Although the APP-null mice are fertile and the mutant mice can be obtained by breeding the homozygous null mice, it is advisable to perform heterozygous intercrosses to obtain littermates as controls. Because genetic backgrounds can influence the phenotype under study, to minimize the variability caused by the genetic background, it is recommended to back-cross the mutant mice onto the C57BL/6 strain for at least 5 and preferably 10 generations. In fact, the APP-null mice available from Jackson Laboratories (Bar Harbor, ME) have been back-crossed for more than 10 generations and are considered congenic. It is particularly important to backcross the animals if a series of behavioral evaluations are planned.

ACKNOWLEDGMENTS The author is indebted to Allan Bradley, who fostered an ideal training and research environment. I also wish to thank many of my colleagues at Merck Research Laboratories who contributed to the generation of APP knockout mice.

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a.

+/+

+/-

+/+

+/-

-/-

-/-

285 APP

185

β-actin

+/+

b.

+/+

-/-

-/116

APP 97.4

β-actin

FIGURE 14.4 APP expression analysis. (a) Top: Northern blotting of total brain RNA from wild-type (+/+), heterozygous (+/–), and homozygous (–/–) APP mice using full-length APP695 cDNA as a probe. Bottom: Mouse β-actin cDNA hybridization for loading control. (b) Top: Western blot analysis of brain APP protein using an APP C-terminal antibody. Bottom: Anti-β-actin antibody staining for loading control.

REFERENCES 1. Goate, A. et al. Segregation of a missense mutation in the amyloid precursor protein gene with familial Alzheimer’s disease, Nature, 349, 704, 1991. 2. Sherrington, R. et al. Cloning of a gene bearing missense mutations in early-onset familial Alzheimer’s disease, Nature, 375, 754, 1995. 3. Levy-Lahad, E. et al. Candidate gene for the chromosome 1 familial Alzheimer’s disease locus, Science, 269, 973, 1995. 4. Yoshikai, S. et al. Genomic organization of the human amyloid beta-protein precursor gene [published erratum appears in Gene, June 30, 1991, 102, 291], Gene, 87, 257, 1990.

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5. Lamb, B.T. et al. Introduction and expression of the 400-kilobase amyloid precursor protein gene in transgenic mice [published erratum appears in Nat. Genet. Nov. 1993, 5, 312], Nat. Genet., 5, 22, 1993. 6. Buxbaum, J.D. et al. Evidence that tumor necrosis factor alpha converting enzyme is involved in regulated alpha-secretase cleavage of the Alzheimer amyloid protein precursor, J. Biol. Chem., 273, 27765, 1998. 7. Lammich, S. et al. Constitutive and regulated alpha-secretase cleavage of Alzheimer’s amyloid precursor protein by a disintegrin metalloprotease, Proc. Natl. Acad. Sci. USA, 96, 3922, 1999. 8. Vassar, R., The beta-secretase, BACE: a prime drug target for Alzheimer’s disease, J. Mol. Neurosci., 17, 157, 2001. 9. Annaert, W. and De Strooper, B., A cell biological perspective on Alzheimer’s disease, Annu. Rev. Cell Dev. Biol., 18, 25, 2002. 10. Moehlmann, T. et al. Presenilin-1 mutations of leucine 166 equally affect the generation of the Notch and APP intracellular domains independent of their effect on Abeta 42 production, Proc. Natl. Acad. Sci. USA, 99, 8025, 2002. 11. Caporaso, G.L. et al. Morphologic and biochemical analysis of the intracellular trafficking of the Alzheimer beta/A4 amyloid precursor protein, J. Neurosci., 14 (Pt 2), 3122, 1994. 12. Koo, E.H. et al. Precursor of amyloid protein in Alzheimer disease undergoes fast anterograde axonal transport, Proc. Natl. Acad. Sci. USA, 87, 1561, 1990. 13. Sisodia, S.S. et al. Identification and transport of full-length amyloid precursor proteins in rat peripheral nervous system, J. Neurosci., 13, 3136, 1993. 14. Yamazaki, T., Selkoe, D.J., and Koo, E.H., Trafficking of cell surface beta-amyloid precursor protein: retrograde and transcytotic transport in cultured neurons, J. Cell Biol., 129, 431, 1995. 15. Schubert, W. et al. Localization of Alzheimer beta A4 amyloid precursor protein at central and peripheral synaptic sites, Brain Res., 563, 184, 1991. 16. Shigematsu, K., McGeer, P.L., and McGeer, E.G., Localization of amyloid precursor protein in selective postsynaptic densities of rat cortical neurons, Brain Res., 592, 353, 1992. 17. Akaaboune, M. et al. Developmental regulation of amyloid precursor protein at the neuromuscular junction in mouse skeletal muscle, Mol. Cell Neurosci., 15, 355, 2000. 18. Jin, L.W. et al. Peptides containing the RERMS sequence of amyloid beta/A4 protein precursor bind cell surface and promote neurite extension, J. Neurosci., 14, 5461, 1994. 19. Morimoto, T. et al. Involvement of amyloid precursor protein in functional synapse formation in cultured hippocampal neurons, J. Neurosci. Res., 51, 185, 1998. 20. Ohsawa, I. et al. Amino-terminal region of secreted form of amyloid precursor protein stimulates proliferation of neural stem cells, Eur. J. Neurosci., 11, 1907, 1999. 21. Rossjohn, J. et al. Crystal structure of the N-terminal, growth factor-like domain of Alzheimer amyloid precursor protein, Nat. Struct. Biol., 6, 327, 1999. 22. Gunawardena, S. and Goldstein, L.S., Disruption of axonal transport and neuronal viability by amyloid precursor protein mutations in Drosophila, Neuron, 32, 389, 2001. 23. Kamal, A. et al. Axonal transport of amyloid precursor protein is mediated by direct binding to the kinesin light chain subunit of kinesin-I, Neuron, 28, 449, 2000. 24. Kamal, A. et al. Kinesin-mediated axonal transport of a membrane compartment containing beta-secretase and presenilin-1 requires APP, Nature, 414, 643, 2001.

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25. Cupers, P. et al. The amyloid precursor protein (APP) cytoplasmic fragment generated by gamma-secretase is rapidly degraded but distributes partially in a nuclear fraction of neurones in culture, J. Neurochem., 78, 1168, 2001. 26. Kimberly, W.T. et al. The intracellular domain of the beta-amyloid precursor protein is stabilized by Fe65 and translocates to the nucleus in a notch-like manner, J. Biol. Chem., 276, 40288, 2001. 27. Cao, X. and Sudhof T.C., A transcriptionally active complex of APP with Fe65 and histone acetyltransferase Tip60, Science, 293, 115, 2001. 28. Kopan, R. and Goate A., A common enzyme connects notch signaling and Alzheimer’s disease, Genes Dev., 14, 2799, 2000. 29. Baek, S.H. et al. Exchange of N-CoR corepressor and Tip60 coactivator complexes links gene expression by NF-kappaB and beta-amyloid precursor protein, Cell, 110, 55, 2002. 30. Leissring, M.A. et al. A physiologic signaling role for the gamma -secretase-derived intracellular fragment of APP, Proc. Natl. Acad. Sci. USA, 99, 4697, 2002. 31. Gao, Y. and Pimplikar, S.W., The gamma -secretase-cleaved C-terminal fragment of amyloid precursor protein mediates signaling to the nucleus, Proc. Natl. Acad. Sci. USA, 98, 14979, 2001. 32. Hardy, J., New insights into the genetics of Alzheimer’s disease, Ann. Med., 28, 255, 1996. 33. Suzuki, N. et al. An increased percentage of long amyloid beta protein secreted by familial amyloid beta protein precursor (beta APP717) mutants, Science, 264, 1336, 1994. 34. Cai, X.D., Golde, T.E., and Younkin, S.G., Release of excess amyloid beta protein from a mutant amyloid beta protein precursor, Science, 259, 514, 1993. 35. De Strooper, B. et al. Deficiency of presenilin-1 inhibits the normal cleavage of amyloid precursor protein, Nature, 391, 387, 1998. 36. Herreman, A. et al. Total inactivation of gamma-secretase activity in presenilindeficient embryonic stem cells, Nat. Cell Biol., 2, 461, 2000. 37. Zhang, Z. et al. Presenilins are required for gamma-secretase cleavage of beta-APP and transmembrane cleavage of Notch-1, Nat. Cell Biol., 2, 463, 2000. 38. Selkoe, D.J., The cell biology of beta-amyloid precursor protein and presenilin in Alzheimer’s disease, Trends Cell Biol., 8, 447, 1998. 39. Zheng, H. et al., Beta-amyloid precursor protein-deficient mice show reactive gliosis and decreased locomotor activity, Cell, 81, 525, 1995. 40. Bradley, A. et al. Formation of germ-line chimaeras from embryo-derived teratocarcinoma cell lines, Nature, 309, 255, 1984. 41. Robertson, E. et al. Germ-line transmission of genes introduced into cultured pluripotential cells by retroviral vector, Nature, 323, 445, 1986. 42. Smithies, O. et al. Insertion of DNA sequences into the human chromosomal betaglobin locus by homologous recombination, Nature, 317, 230, 1985. 43. Thomas, K.R. and Capecchi M.R., Site-directed mutagenesis by gene targeting in mouse embryo-derived stem cells, Cell, 51, 503, 1987. 44. Mills, A.A. and Bradley, A., From mouse to man: generating megabase chromosome rearrangements, Trends Genet., 17, 331, 2001. 45. Bradley, A. et al. Modifying the mouse: design and desire, Biotechnology, 10, 534, 1992. 46. Bradley, A. et al. Genetic manipulation of the mouse via gene targeting in embryonic stem cells, Ciba Found. Symp., 165, 256, 1992. 47. Capecchi, M.R., Targeted gene replacement, Sci. Am., 270, 52, 1994.

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48. Capecchi, M.R., Generating mice with targeted mutations, Nat. Med., 7, 1086, 2001. 49. Mansour, S.L., Thomas, K.R., and Capecchi, M.R., Disruption of the proto-oncogene int-2 in mouse embryo-derived stem cells: a general strategy for targeting mutations to non-selectable genes, Nature, 336, 348, 1998. 50. Izumi, R. et al. Positive and negative regulatory elements for the expression of the Alzheimer’s disease amyloid precursor-encoding gene in mouse, Gene, 112, 189, 1992. 51. Bradley, A., Zheng, B., and Liu, P., Thirteen years of manipulating the mouse genome: a personal history, Int. J. Dev. Biol., 42, 943, 1998. 52. Robertson, E.J., Teratocarcinomas and Embryonic Stem Cells: A Practical Approach: Embryo-Derived Stem Cell Lines, IRL Press, Oxford, 1987, ch. 4 and ch. 5. 53. Ramirez-Solis, R. et al. Genomic DNA microextraction: a method to screen numerous samples, Anal. Biochem., 201, 331, 1992. 54. Wong, P.C., et al. Presenilin 1 is required for Notch1 and DII1 expression in the paraxial mesoderm, Nature, 387, 288, 1997. 55. Dawson, G.R. et al. Age-related cognitive deficits, impaired long-term potentiation and reduction in synaptic marker density in mice lacking the beta-amyloid precursor protein, Neuroscience, 90, 1, 1999. 56. Phinney, A.L. et al. Aged APP-null mice exhibit a learning impairment which is not mediated by a loss of hippocampal neuron or synaptic bouton number, Neuroscience, 90, 1207, 1999. 57. Seabrook, G.R. et al. Mechanisms contributing to the deficits in hippocampal synaptic plasticity in mice lacking amyloid precursor protein, Neuropharmacology, 38, 349, 1999.

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a

b TGN Membrane 0’

15’ 30’

60’ 90 ’

TGN Vesicles

ER Membrane

0’ 15 ’ 30 ’ 60’ 90 ’

PS1WT

PS1WT

PS1 -/-

PS1 -/-

6E10

c

FITC -VVA

ER Vesicles

0’ 15’ 30’ 60 ’ 90 ’

0’ 15 ’ 30’ 60’ 90 ’

d Newly Synthesized βAPP on Cell Surface

Total Cell βAPP WT WT

∆Μ1,2 ∆Μ1,2 Chase Time (min)

0’

10’

20’

30 ’

45’

60 ’ 120 ’

COLOR FIGURE 6.4 PS1 deficiency or loss of function accelerates βAPP trafficking from the TGN/ER and increases cell surface delivery.

COLOR FIGURE 8.2 Intracellular accumulation of aggregated Aβ42 in DS astrocytes.

© 2005 by CRC Press LLC

COLOR FIGURE 8.3 Double labeling of Aβ42 and different subcellular markers.

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COLOR FIGURE 8.4 Mitochondrial dysfunction in normal astrocytes induces aberrant APP processing and intracellular Aβ accumulation.

COLOR FIGURE 8.5 Impaired mitochondrial function in DS astrocytes.

© 2005 by CRC Press LLC

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a. Aß injection

AT100

Aß injection+LiCl

AT100

b.

Vehicle Aß

Aß+LiCl

AT180

Vehicle Aß Aß+LiCl

Tau-C

SDS-insoluble tau COLOR FIGURE 10.3 The effects of GSK-3β inhibition on the Aβ-induced formation of NFT-like tau pathology.

© 2005 by CRC Press LLC

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COLOR FIGURE 12.1 Detection of fibrillar amyloid by thioflavin S and Congo red staining.

COLOR FIGURE 12.8 Double immunofluorescent labeling of plaques (R1282) and microglia (CD45) showed similar patterns of colocalization in Aβ immunized (bottom) and untreated (top and middle) PSAPP mice after 8 weeks of Aβ immunization.

© 2005 by CRC Press LLC